An Atlas of Human Blastocysts (Encyclopedia of Visual Medicine Series)

An Atlas of Human Blastocysts

THE ENCYCLOPEDIA OF VISUAL MEDICINE SERIES

An Atlas of Human Blastocysts Lucinda L.Veeck, MLT, hDSC Assistant Professor of Embryology in Obstetrics and Gynecology Assistant Professor of Reproductive Medicine Director, Embryology Laboratories The Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University, New York and Nikica Zaninović, MS Supervisor, Embryology Laboratories The Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University, New York

The Parthenon Publishing Group International Publishers in Medicine, Science & Technology A CRC PRESS COMPANY BOCA RATON LONDON NEW YORK WASHINGTON, D.C.

Cover photo: Human blastocyst stained with fluorescence dyes: Bcl-2-FITC (green), DAPI (blue) ‘Thumbnail’ figures: Human frozen-thawed single-pronucleate conceptus donated for time-lapse studies: cleavage from the eight-cell stage of development through to the hatched blastocyst stage Published in the USA by The Parthenon Publishing Group 345 Park Avenue South, 10th Floor New York, NY 10010 USA Published in the UK and Europe by The Parthenon Publishing Group 23–25 Blades Court Deodar Road London SW15 2NU UK Copyright © 2003 The Parthenon Publishing Group Library of Congress Cataloging-in-Publication Data An atlas of human blastocysts/[edited by] Lucinda L.Veeck and Nikica Zaninovic. p. ; cm.—(The encyclopedia of visual medicine series) Includes bibliographical references and index. ISBN 1-84214-169-4 (alk. paper) 1. Fertilization in vitro, Human—Atlases. 2. Blastocyst—Atlases. 3. Human reproductive technology—Atlases. I.Veeck, Lucinda L. II. Zaninovic, Nikica. III. Series. [DNLM: 1. Blastocyst—cytology—Atlases. 2. Preimplantation Phase—physiology—Atlases. 3. Fertilization in Vitro—Atlases. WQ 17 A8808 2003] RG135A876 2003 618.1′78059–dc21 2003040564 British Library Cataloguing in Publication Data Veeck, Lucinda L. An atlas of human blastocysts 1. Blastocyst—Atlases 2. Human reproductive technology— Atlases I. Title II. Zaninovic, Nikica 618.1'78059 ISBN 0-203-00893-6 Master e-book ISBN

ISBN 1-84214-169-4 (Print Edition) First published in 2003 This edition published in the Taylor & Francis e-Library, 2005. "To purchase your own copy of this or any of Taylor & Francis or Routledge’s collection of thousands of eBooks please go to http://www.ebookstore.tandf.co.uk/." No part of this book may be reproduced in any form without permission from the publishers except for the quotation of brief passages for the purposes of review Composition by The Parthenon Publishing Group

Contents List of contributing authors

vi

Foreword

ix

Preface

x

Acknowledgements

1. Overview of early human preimplantation development in vitro

xiii

1

2. Metabolic requirements during preimplantation development and the formulation of culture media David K.Gardner and Michelle Lane 3. Human morulae in vitro

37

4. Cell allocation and differentiation

97

66

5. Human blastocysts in vitro

123

6. Preembryo selection and blastocyst quality: how to choose the optimal conceptus for transfer 7. Blastocyst hatching

182

8. Blastocyst cryopreservation and thawing

231

9. Cell death (apoptosis) In human blastocysts Kate Hardy, Sophie Spanos and David L.Becker 10. Human implantation Owen Davis and Zev Rosenwaks 11. Human embryonic stem cells Michal Amit and Joseph Itskovitz-Eldor 12. The mammalian blastocyst as an experimental model Shoukhrat M.Mitalipov, Hung-Chih Kuo and Don P.Wolf 13. The moral status of the human blastocyst Howard W.Jones, Jr

210

250 277 288 312 339

Glossary of terms

350

Abbreviations and symbols used for embryology documentation

367

Index

370

List of contributing authors Michal Amit Department of Obstetrics and Gynecology Rambam Medical Center PO Box 9602 Haifa 31096 Israel David L.Becker Department of Anatomy and Developmental Biology University College London Gower Street London WC1 6BT UK Owen Davis Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University New York USA Kate Hardy Institute of Reproductive and Developmental Biology Imperial College Hammersmith Hospital Du Cane Road London W12 0NN UK Joseph Itskovitz-Eldor Department of Obstetrics and Gynecology Rambam Medical Center PO Box 9602 Haifa 31096 Israel David K.Gardner Colorado Center for Reproductive Medicine Englewood Colorado 80110 USA

Howard W.Jones, Jr Jones Institute for Reproductive Medicine 610 Colley Avenue Norfolk Virginia 23507 USA Hung-Chih Kuo Oregon National Primate Research Center 505 NW 185th Avenue Beaverton Oregon 97006 USA Michelle Lane Colorado Center for Reproductive Medicine Englewood Colorado 80110 USA Shoukhrat M.Mitalipov Oregon National Primate Research Center 505 NW 185th Avenue Beaverton Oregon 97006 USA Zev Rosenwaks Center for Reproductive Medicine and Infertility Weill Medical College of Cornell University New York USA Sophie Spanos Institute of Reproductive and Developmental Biology Imperial College Hammersmith Hospital Du Cane Road London W12 0NN UK Don P.Wolf Oregon National Primate Research Center 505 NW 185th Avenue Beaverton Oregon 97006

and Departments of Obstetrics and Gynecology, and Physiology and Pharmacology Oregon Health and Science University Portland Oregon 97201 USA

Foreword Assisted reproductive technology has many intriguing aspects, but two are of surpassing fascination. First is the breathlessness of clinical result. With success, there is elation and presumed understanding of the physiological process; with failure, there is a frustration that leads the scientific community to strive for better understanding and more satisfactory results. Second is the moving experience of viewing microscopically the morphological processes that lead to an independent being—in this case a human being. The enigma is to distinguish the potential for viability from the many observed variations, which, according to our present understanding, are often expressions of the genetic misfits and misfires that are characteristic of human reproduction. Are we having a glimpse of the evolutionary process? From its microscopic origin, the preembryo blooms into the human form, a process both wondrous and scientifically intriguing. Heeding biological commands, cells grow purposefully according to a predefined plan, endowing the new genetic entity with viability and function. Now, there is even more; as our experience extends beyond morphology. We are beginning to understand something about genes, gene products, enzymes, and proteins which drive these morphological characteristics. The excitement and reality of clinical and laboratory work are captured by Lucinda Veeck, Nikica Zaninović, and their collaborators in An Atlas of Human Blastocysts. This is a book for those who wish to experience the satisfaction of being certain that they are up to date regarding extended culture procedures and the complexities of blastocyst development, considered key to achieving high pregnancy rates while minimizing the troublesome complication of multiple pregnancies. In the following pages, the reader is given an opportunity to study the human, rhesus and murine blastocyst under optimal clinical and research conditions. The blastocyst’s nutrient requirements during culture, its growth through various key stages, and its ability to survive freezing and thawing are all examined. We are guided through the early aspects of cell allocation and differentiation and are enlightened to the processes of hatching and programmed cell death. In sum, the reproductive process is demonstrated photographically from fertilization through to completion of implantation. Additionally, and of great interest, current scientific research applications are included. Leaders in various clinical and scientific fields have come together to create this superb volume. An Atlas of Human Blastocysts is a dynamic and authoritative collection of microanatomical examples and definitively captures the earliest events of mammalian development in vitro. It is an absolute ‘must-read’ for clinicians and scientists working in the field of assisted reproduction. Howard W.Jones, Jr, MD Georgeanna Seegar Jones, MD

Preface Why have those of us working in assisted reproductive laboratories become so suddenly fascinated with blastocysts? The answers are simple. First and foremost, never before in history have we had the opportunity to study closely human blastocyst development in vitro. Early descriptions of human morulae and blastocysts often relied on studying discarded material grown under suboptimal culture conditions after in vitro fertilization (IVF) trials. Investigating morphology, growth rate, metabolic requirements and genetic factors under these conditions probably led us to many misleading conclusions. Only with the development of sequential media have we been able routinely to grow viable blastocysts in our laboratories with some measure of confidence. Without doubt, in vitro culture techniques will continue to improve as additional knowledge is gained, enabling us to understand better the human reproductive process and ultimately provide our patients with tremendous benefit. Second, we recognize that, through in vitro developmental investigations involving extended culture, we have been given the opportunity to offer a much improved and safer service to our patients by reducing the number of preembryos for transfer. How often in the past have we observed patients desperately desiring a healthy child, anxious to receive three or more pre-embryos for transfer, and then watched them agonize guiltily when forced to reduce selectively a high-order multiple pregnancy? This sad treatment option is all too often necessary because higher-order gestations, those involving more than two fetal hearts on ultrasound examination, are the largest single cause of poor obstetric outcome and subsequent neonatal difficulties. Triplet and quadruplet pregnancies are associated with high incidences of preeclampsia, gestational diabetes, pregnancy-induced, hypertension, preterm labor, low birth weight and extensive neonatal care1. Although multifetal pregnancy reduction to twins is an option, the procedure itself carries medical and emotional risks2,3. Clearly, the most efficient way to avoid any form of multiple pregnancy is to limit the number of preembryos for intrauterine transfer to a single conceptus. While straightforward in theory, the reality of this approach leaves much to be desired. Indeed, most IVF programs experience no greater than a 20–30% clinical pregnancy rate per transfer when a single 4–8-cell conceptus is replaced. With treatment costs of $5000– $15000 per IVF attempt in the United States, often not covered by insurance, this figure is too low to be cost-effective or desirable to the couple being treated. For this reason, more than one, and frequently more than three, day-2 or day-3 preembryos have been routinely replaced in an effort to optimize the chances for pregnancy. Therein lies the problem: multiple transfer of early developing preembryos carries the risk of plural gestation, a risk that, until recently, could not be fully eliminated without decreasing the overall likelihood of pregnancy. In the Cornell program, one in three young women under the age of 34 years will establish a multiple pregnancy if three preembryos are replaced on day 3, and 20% of women aged 34–39 years old will follow the same pattern. Because this trend is seen world-wide, it has become the recommended policy of many IVF

centers to replace no more than two conceptuses whenever possible, many countries mandating this by law. The incidence of multiple pregnancy has risen throughout the world as a consequence of assisted reproductive technologies. It has been reported that the rate of triplet and higher-order gestation infants per 100 000 Caucasian live births in the United States increased by 191 % between 1972 and 1991, with 38% due to assisted conceptions and 30% to increased child-bearing among older women4. Another negative aspect to the rising rate of multiple pregnancy involves the associated economic burden resulting from preterm birth and increased hospital stays. Analysis of births at Brigham and Women’s Hospital in Boston between 1986 and 1991 revealed that assisted reproductive technologies accounted for 2% of single, 35% of twin and 77% of triplet deliveries in that particular unit. Hospital charges per single baby averaged $9845, for twins they averaged $37947 ($18974 per baby) and for triplets they averaged $109765 ($36588 per baby)5. Since these figures are more than a decade old, one may assume that hospital costs are even higher today for couples experiencing multiple births. Unfortunately, early demise of the human conceptus is a common event. Approximately 73% of natural single conceptions are lost before reaching 6 weeks of gestation, and, of the remainder, roughly 90% survive to term6. Although conceptions from IVF do nearly as well as natural pregnancies after clinical recognition, they result in higher losses between the onset of fertilization and completion of implantation, presumably due to developmental arrest or unrecognized abnormalities. Realization of this shortcoming prompts patients to ask for the replacement of multiple preembryos and allows us to agree in an effort to optimize ongoing pregnancy rates. Nevertheless, the necessity of replacing more than a single preembryo in order to establish good pregnancy rates would be moot if one could appropriately choose for transfer the healthiest and most viable conceptus from a cohort of growing preembryos. Imagine one day in the future when our patients will receive a single, healthy hatched blastocyst while having all other potentially viable ones frozen. The incidence of twin and greater gestations would be effectively eradicated! It is not only reasonable, but prudent, to enquire which factors contribute to preembryo viability. Certainly, genetic stability is a major prerequisite for the implantation and delivery of a healthy child. At present, we know little about the genetic make-up of the preembryos within our incubators unless they are biopsied and examined, hardly a practical screening modality for every preembryo growing in the laboratory. Yet, when these examinations are performed, evidence comes to light that chromosomal abnormalities, both numerical and structural, are often associated with fragmented, multinucleated or poorly developing preembryos, and, conversely, many preembryos presenting good morphology possess lethal genetic aberrations. This leads us to recognize that, although morphological evaluations may furnish clues that minimally enhance our proficiency at choosing the best preembryos for transfer, these systems are severely limited in their ability to provide rock-hard evidence for subsequent normal development. Only by using new and very exciting non-invasive methods of assessment, such as amino acid profiling, will we enter a new era for diagnostic preembryo selection7. In further support of blastocyst transfer, it has been observed that genetically unhealthy preembryos often cease growth at very early cleavage stages. Almeida and Bolton proposed that there is a progressive loss of chromosomally abnormal preembryos

after pronuclear development to at least the 8-cell stage8. When preembryos of varying morphological grades were studied cytogenetically, these investigators found a 65% incidence of abnormality at the pronuclear stage, a 55% incidence at the 2–4-cell stage and a 27% incidence at the 5–8-cell stage. Preembryos with poor morphology demonstrated almost a three-fold increase in chromosomal anomalies as compared to those with good morphology. From these data, it is logical to deduce that some form of natural selection continues beyond the 8-cell stage, perhaps through early fetal development. Might we not expect progressive natural selection to occur if we extend culture times beyond the standard 2 or 3 days? Will blastocyst transfer allow us successfully to replace a single conceptus? The purpose of the following text and photographic collection is to demonstrate to the reader that extended culture to blastocyst stages of development is now indeed an achievable option in our laboratories. Lucinda L.Veeck Nikica Zaninović ‘Faith’ is a fine invention when Gentlemen can see. But Microscopes are prudent in an emergency Emily Dickinson (contributed by Helen Maloney)

It will be found that everything depends on the composition of the forces with which these particles of matter act upon one another: and from these forces, as a matter of fact, all phenomena of Nature take their origin Ru er Bošković, Croatian scientist (The Theory of Natural Philosophy, 1758)

References 1. Skupski DW, Nelson S, Kowalik A, et al. Multiple gestations from in vitro fertilization: successful implantation alone is not associated with subsequent preeclampsia. Am J Obstet Gynecol 1996; 175:1029–32 2. Melgar CA, Rosenfeld DL, Rawlinson K, Greenberg M. Perinatal outcome after multifetal reduction to twins compared with nonreduced multiple gestations. Obstet Gynecol 1991; 78:763–7 3. Groutz A, Yovel I, Amit A, Yaron Y, Azem F, Lessing JB. Pregnancy outcome after multifetal pregnancy reduction to twins compared with spontaneously conceived twins. Hum Reprod 1996; 11:1334–6 4. Wilcox LS, Kiely JL, Melvin CL, Martin MC. Assisted reproductive technologies: estimates of their contribution to multiple births and newborn hospital days in the United States. Fertil Steril 1996; 65:361–6 5. Callahan TL, Hall JE, Ettner SL, Christiansen CL, Greene MF, Crowley WF Jr. The economic impact of multiplegestation pregnancies and the contribution of assisted-reproduction techniques to their incidence. N Engl J Med 1994; 331:244–9 6. Boklage CE. Survival probability of human conceptions from fertilization to term. Int J Fertil 1990; 35:75, 79–80, 81–94 7. Houghton FD, Hawkhead JA, Humpherson PG, et al. Non-invasive amino acid turnover predicts human embryo developmental capacity. Hum Reprod 2002; 17:999–1005 8. Almeida PA, Bolton VN. The relationship between chro mosomal abnormality in the human preimplantation embryo and development in vitro. Reprod Fertil Dev 1996:8:235–41

Acknowledgements Sincere appreciation is extended to the physicians, nurses and support staff of the Institute for Reproductive Medicine of Weill Medical College of Cornell University/New York Hospital for their direct or indirect contribution to the work detailed in this book. Very special acknowledgement is given to the many hard-working embryologists and laboratory support staff at Cornell who make up a quite dedicated and unique team of professionals: Rosemary Berrios, Richard Bodine, Jose Bustamante, Robert Clarke, Carol Ann Cook, Margarita Fienco, June Hariprashad, Myriam Jackson, Deborah Liotta, Rose Moschini, Gianpiero Palermo, Jason Park, Patricia Pascal Roy, Takumi Takeuchi, Kangpu Xu and Zhen Ye; and our sadly missed friends, David Travassos and Eric Urcia. We also thank Dr Michael Bedford and Mrs Pamela Sully of Weill Medical College for editorial assistance, and offer extreme gratitude to those who contributed photographic material or text: Dr Michal Amit, Dr David Becker, Dr Owen Davis, Dr David Gardner, Dr Kate Hardy, Drs Howard and Georgeanna Jones, Dr Hung-Chi Kuo, Dr Joseph Itskovitz-Eldor, Dr Michelle Lane, Dr Shoukhrat Mitalipov, Dr Zev Rosenwaks, Dr Sophie Spanos and Dr Don Wolf. Most figures were photographed using a Nikon Diaphot TE300 microscope equipped with a Sony CatsEye DKC-5000 camera. Parthenon Publishing is responsible for the color matching and placement of selected photographs.

1 Overview of early human preimplantation development in vitro Ovulation induction for assisted reproductive procedures The cornerstone of successful assisted reproductive technology (ART) has been the ability to replace several selected preembryos from a larger cohort obtained following recruitment, harvest and fertilization of multiple oocytes. Thus, although the first successful human in vitro fertilization (IVF) pregnancy followed the retrieval of a single oocyte in a spontaneous menstrual cycle1, current standard practice in ART programs worldwide entails the use of controlled ovarian hyperstimulation in order to maximize pregnancy rates. This strategy, while maximizing pregnancy rates, has also been associated with the inherent increased risks of multiple pregnancies. A wide variety of ovulation-inducing agents have been employed in the practice of ART, including clomiphene citrate, human menopausal gonadotropins (hMG), and recombinant gonadotropin preparations, with and without the adjunctive use of gonadotropin releasing hormone (GnRH) agonists and antagonists. Currently, the dominant approach to ovulation induction for IVF combines exogenous gonadotropins (hMG, Puriffied follicle stimulating hormone (FSH) and recombinant FSH) with GnRH agonists. Although clomiphene citrate was once extensively employed for ART, either as a single agent or in combination with gonadotropins, the current dominance of pure gonadotropin-based protocols was spurred by the premise that this approach is more physiological and might avoid the potentially detrimental effects of clomiphene on the oocytes and endometrium. Commercially available gonadotropin preparations include: hMG, formulated in ampules containing 75 IU each of FSH and luteinizing hormone (LH); purified FSH, containing 75 IU of FSH with less than 1 IU of LH and, more recently, recombinant FSH. Urinary-derived gonadotropins are heterogeneous with respect to glycosylation and the presence of degraded fragments, which can lead to varying biopotency among batches, whereas recombinant gonadotropins are uniform. Over 2000 different GnRH agonists have been synthesized. Endogenous GnRH is rapidly degraded by cleavage at the Gly6-Leu7 and Pro9-Gly10 positions, with a resultant half-life of less than 10 min. The selective substitution of amino acids at positions 6 and 10 of the GnRH molecule leads both to enhanced binding affinity to the GnRH receptor and decreased susceptibility to degradation by endopeptidases, thus prolonging the halflife and enhancing the biological activity of these agents. The pharmacological response to GnRH agonist administration is biphasic, with an initial surge of gonadotropin release from the adenohypophysis, but prolonged GnRH receptor occupancy results in desensitization and down-regulation of the gonadotropes, thus effecting reversible hypogonadism. The adjunctive use of GnRH agonists in IVF has several apparent

An atlas of human blastocysts 2

advantages, including a reduction in the incidence of untimely LH surges and premature luteinization, the ability to program the initiation of stimulations so as to permit a more even distribution of a clinic’s workload, and, most significantly, an overall improvement in IVF success rates, a finding confirmed by a published meta-analysis of randomized, controlled trials2. When applied to IVF, GnRH agonists may be administered either in a long or a short protocol. In the long protocol, currently favored by most centers, GnRH agonist treatment is initiated in the mid-luteal phase of the preceding menstrual cycle; pituitary downregulation ensues within 5–10 days, andis indicated by the onset of menses. Gonadotropin therapy is then undertaken concurrently, typically commencing on cycle day 3 or once adequate suppression of estradiol is documented. The dosage of gonadotropins ranges from two to four ampules per day, with higher doses occasionally employed in patients predicted to have a poor response. The cycle is monitored with daily estradiol determinations commencing after 2–3 days of therapy; serial sonographic follicular measurements are performed once the estradiol exceeds a threshold level, generally by the sixth or seventh day of the cycle. The daily dosage of gonadotropins may be adjusted according to the individual patient’s response, e.g. with a step-down once follicular recruitment has been achieved, in an effort to attain greater synchronization of follicular maturation and a reduced risk for the development of ovarian hyperstimulation syndrome. Appropriate timing of the ovulatory dose of human chorionic gonadotropin (hCG) is critical for the retrieval of an adequate number of optimally mature oocytes, and is determined by parameters including the mean diameter of the lead follicles (typically >16 mm), the absolute estradiol level (e.g. >500 pg/ml) and the pattern of estradiol rise and follicular growth. The GnRH agonist is discontinued on procedure, performed transvaginally with ultrasound the day of hCG administration. The oocyte retrieval guidance, is typically undertaken 34–36 h following the administration of hCG. In the short or ‘flare’ GnRH agonist protocols, the agonist is initiated in the early follicular phase, usually on cycle day 2 or 3. Concurrent therapy with gonadotropins commences 1–3 days later. This approach exploits the agonist phase of GnRH agonist treatment, thus reducing the total dosage requirement for gonadotropins and shortening the duration of stimulation. Although both long and short protocols have their adherents, the former approach is more prevalent. More recently, GnRH antagonists have been introduced, which allow for late follicular suppression of the LH surge, eliminating the need for prolonged pretreatment down-regulation. The goal of all ovulation induction protocols for ART is to permit the recruitment and harvest of an optimal number of preovulatory oocytes, so as to maximize clinical efficiency. Pregnancy rates may thus be optimized through the selection and transfer of a few of the ‘best quality’ preembryos, with the option of cryopreserving potentially viable conceptuses in excess of that number.

Gametes In most species, there are just two types of gametes, and they are radically different. Apart from motor neurons with their remarkably long axons, the oocyte is among the

Overview of early human preimplantation Overview of early human preimplantation

3

largest cells of the human organism. Conversely, spermatozoa and red blood cells are two of the smallest. The diameter of the mature human oocyte is approximately 110–115 µm, and it is bounded by a plasma membrane called the oolemma. Surrounding the oocyte/oolemma is a glycoprotein envelope called the zona pellucida, a structure approximately 15–20 µm wide (becoming a bit thinner after fertilization) that protects the oocyte during transport and fertilization. Between the oolemma and the zona pellucida is the fluid-filled perivitelline space. The use of this term persists despite its inaccuracy when describing the oocytes of humans or most other mammals; it acknowledges the word vitellus, a term traditionally used to describe the yolky substance of a hen’s egg, which contains abundant nutrient reserves. The cytoplasm of the mammalian oocyte is usually referred to as the ooplasm, a more appropriate term for describing the living portion of the human gamete. The main organelles of the ooplasm are the mitochondria, the endoplasmic reticulum and the Golgi system. When fully capable of undergoing a normal fertilization process, the secondary oocyte is briefly arrested in its course of maturation at metaphase II of meiosis. Nuclear maturation is usually closely attended by a general maturation of the cytoplasm, and is characterized by an increase in the number of organelles scattered throughout the ooplasm. The presence of a first polar body conveys that nuclear maturation has reached this stage. Along with the zona pellucida and perivitelline space, the total diameter of the mature human oocyte is approximately 150 µm. An oocyte incubated with spermatozoa before reaching metaphase II may incorporate a spermatozoon into its ooplasm and yet fail to initiate events leading to sperm decondensation; such an oocyte ultimately lacks a functional male pronucleus3. One study examining 518 non-fertilized oocytes demonstrated that 22% had actually been penetrated by sperm, but without oocyte activation or pronuclear formation4. Many of these oocytes may have been immature when combined with spermatozoa. Besides the requirement for nuclear maturation, it is believed that a brief period is necessary after extrusion of the first polar body for the oocyte to gain full cytoplasmic competence. An oocyte that is meiotically mature but slightly underdeveloped or overdeveloped with regard to its cytoplasm may be more apt to display one, three or more pronuclei. With immature cytoplasm, the cortical granule numbers and response may be inadequate; with postmature cytoplasm, cortical granule release may be inhibited owing to the inward migration of the granules towards the interior of the cell. In either instance, there is evidence that the zona reaction is also often poorly functional when the spermoocyte interaction is not appropriately timed with regard to oocyte nuclear and cytoplasmic maturity5. Oocytes collected for IVF are generally surrounded by several layers of cells, which define the cumulus oophorus. Cells of the cumulus are instrumental, via gap junctions, in nurturing the oocyte during growth and possibly in passing inhibiting factors (e.g. cyclic adenosine monophosphate (cAMP)) necessary for deterring the resumption of meiosis6. The innermost layer of cells is called the corona or coronal layer. This layer expands and presents a radiant pattern as oocytes mature in response to exogenous hCG or a mid-cycle surge of LH. Near ovulation, as they loosen and expand, cumulus cells are observed to retract from the zona pellucida of the oocyte, presumably cutting off the previously important cellular-oocyte communication. It has been proposed that oocytes not

An atlas of human blastocysts 4

associated with proliferative cellular changes near ovulation have very limited potential for implantation, despite fertilization and apparently normal development in vitro7. In most mammalian species studied in vivo, the oocyte arrives at the site of fertilization in the ampulla of the Fallopian tube still surrounded by the cumulus mass. The cumulus may play a role in assisting transport of the oocyte into the Fallopian tube through fimbrial cilia-cumulus cell contact. Another possible use of the cumulus after oocyte maturation is that its radially arranged cells help to guide spermatozoa towards the oocyte just before fertilization; however, there is no hard evidence for this speculation. Break-up of the cumulus mass is brought about by dissolution of its mucoid hyaluronic acid matrix by enzymes released by the spermatozoa. Follicular membrana granulosa cells disassociated from cumulus cells are found in follicular aspirates collected for IVF. The number of cells collected will vary from follicle to follicle according to the extent of negative pressure exerted during suction, the size of the needle and the overall maturity of the follicle. As with cumulus cells, thecorrelation between morphological aspects of free granulosa cells and oocyte nuclear maturity is not exact, but mature-appearing cells (large, well-dispersed cells) are generally collected along with mature oocytes, and immature-appearing cells (smaller, tightly packed cells) along with immature oocytes. Follicular membrana granulosa cells may be assessed at the time of oocyte harvest to aid in the evaluation of follicular maturity. They are subsequently often used during in vitro studies to examine metabolic activity or steroid synthesis. The oocyte observed while its chromosomes are at metaphase I of maturation requires some time in culture before attaining full meiotic competence8. More than 98% of these oocytes will complete their journey towards metaphase II and first polar body extrusion. Oocytes with chromosomes at prophase I of maturation are truly immature; more than 80% of these will continue through metaphase I to metaphase II if isolated and incubated in an appropriate medium for 24 h. Assessment of maturity Traditionally, evaluation of oocyte maturity has been based upon the expansion and radiance of the cumulus-corona complex which surrounds the harvested oocyte9,10. With this assessment, oocytes are rapidly categorized as mature (correlated to metaphase II of maturation) when they possess an expanded and luteinized cumulus matrix and a radiant or sun-burst corona radiata. A less-expanded cumulus-corona complex denotes an intermediate stage of maturity (correlated to metaphase I of maturation), and absence of expanded cumulus is generally associated with immaturity (correlated to prophase I of maturation). While this type of analysis usually closely approximates the true nuclear status of the oocyte, it is too often imprecise, and may lead to subsequent laboratory errors in the handling of gametes. In fact, nuclear maturation of the oocyte and cellular maturation of the cumulus are frequently disparate11,15. When disparity occurs, immature oocytes may be inseminated prematurely, and fail to produce a favorable outcome. As well as fertilization failure, other detrimental side-effects accompany combining sperm and eggs at suboptimal times; ovulation-induction protocols may not be suitably appraised and male factor issues become difficult to interpret, based on poor fertilization results.

Overview of early human preimplantation Overview of early human preimplantation

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Because of these pitfalls, techniques have been developed to assess more accurately the meiotic status of the oocyte. A systematic approach can be used to produce a maturation score by grading the size of the follicle, expansion of the cumulus mass, radiance of the corona cells, size/cohesiveness of associated membrana granulosa cells and shape/color of the oocyte itself, if visible within the mass of surrounding cellular investments. Alternatively, frank visualization of the oocyte and its germinal vesicle or first polar body can be attempted by spreading out the cumulus mass, or by removing it altogether with the aid of enzymes. If clearly visible or denuded of cells, oocytes are classified according to the presence or absence of first polar bodies/germinal vesicles, and are inseminated/injected accordingly: Metaphase II (MII) First polar body present, no germinal vesicle; inseminated or injected 3–5 h after collection; Metaphase I (MI) No first polar body, no germinal vesicle; inseminated or injected 1– 5 h after extrusion of the first polar body; Prophase I (PI) Germinal vesicle present; inseminated or injected 26–29 h after collection. Our experience has been that oocytes collected at more advanced stages of in vivo maturation demonstrate the greatest ability to form two pronuclei after insemination8,9,11. Fertilization rates drop only slightly when oocytes require a period of 5–15 h in culture before extruding the first polar body, but fertilization is markedly reduced when more than 15 h pass before the maturational process is completed. The reason for this is probably related to sperm functionality as well as oocyte maturity, since processed sperm may be more than 24 h old before being placed with an early MI or PI oocyte. Under these conditions, the precise cause of the lower incidence of fertilization of very immature oocytes is difficult to interpret8. If small follicles are punctured, approximately 20–30% of oocytes collected for IVF are meiotically immature at the time of harvest from the ovary. This is undoubtedly due to the stimulation of multiple follicles during clinical ovulation induction, some large and well-vascularized, and some small with late recruitment. If all oocytes are placed with sperm at the same time, a proportion slightly higher than this percentage will fail to become fertilized normally. Logically enough, when oocytes are placed with sperm only as they have reached full maturity, far better fertilization results are attained. The incidence of abnormal fertilization (one pronucleus, three or more pronuclei) is not different between MII oocytes and MI or PI oocytes that have matured in culture before insemination or injection8,10. Pregnancy potential after the transfer of preembryos developed from MII and MI oocytes is similar, regardless of whether 0 or 20 h has been required for maturation before insemination or injection16. Only preembryos developing from PI oocytes demonstrate a significantly reduced potential for implantation and live birth, although such births are certainly within the realm of possibility17,20. Metaphase II oocyte The MII oocyte (Figure 1.1) is often termed mature, ripened or preovulatory, vague descriptions that fail to specify the exact meiotic status of the gamete. This oocyte is at a resting stage of meiosis II after extrusion of the first polar body and direct passage to

An atlas of human blastocysts 6

MII. Chromosomes are divided between the oocyte and the polar body (23 chromosomes, 46 chromatids, 2n DNA in each), those in the oocyte being attached to spindle microtubules3 (Figure 1.2). For a while after its formation, the first polar body remains connected to the oocyte by the meiotic spindle, forming a cytoplasmic bridge. Chromosomes within the first polar body may remain clumped together, may undergo a second meiotic division or may scatter within the cytoplasm; generally a nucleus is not formed3,21. The first polar body contains cortical granules because of its extrusion before sperm penetration and cortical granule release; in the oocyte, 1–3 layers of cortical granules are present at the periphery. Under the microscope, the oocyte is characterized by its round, even shape and displays an ooplasm of light color and homogeneous granularity. It is usually associated with an expanded, luteinized cumulus and a sun-burst corona radiata. Membrana granulosa cells harvested along with the MII oocyte are loosely aggregated, with mature features8,10,14,20. Metaphase I oocyte The MI oocyte (Figure 1.3) is considered nearly mature or intermediate in maturation. The oocyte has completed prophase of meiosis I; the germinal vesicle and its nucleolus have faded and disappeared. During this stage a spindle forms, and recombined maternal and paternal chromosomes line up randomly towards the poles. Later, at telophase, whole chromosomes sort independently to oocyte or first polar body. An MI oocyte requires 1–24 h in culture before reaching full maturity. Those needing less than 15 h are considered late in maturity, while those requiring more than 15 h are defined as early8–11,14,15. Under the microscope, the MI oocyte is characterized by the absence of both germinal vesicle and first polar body. A late MI oocyte is round and even in form, with homogeneously granular and light-colored ooplasm. Early MI oocytes may display minor central granularity. Mature-appearing cumulus cells are usually associated with late stages. Because first polar body extrusion can occur at any time after harvest, it is necessary to examine the oocyte at regular intervals to determine the correct timing for insemination. If sperm are placed with the oocyte before nuclear and cytoplasmic maturation are complete, they generally fail to decondense within the ooplasm, or abnormal fertilization occurs. If insemination is delayed too long, in vitro aging may follow, with similar undesired consequences3,8 (Figures 1.4 and 1.5). Prophase I oocyte The PI oocyte (Figure 1.6) is often termed immature or unripened. It possesses a tetraploid amount of DNA owing to the presence of 46 double-stranded chromosomes. This oocyte begins to mature in response to gonadotropin surges and reduction in follicular maturation-inhibiting factors. The germinal vesicle, which persisted throughout earlier growth phases, begins its progression to germinal vesicle breakdown (GVBD) and the oocyte enlarges. Most PI oocytes collected for IVF have been stimulated to resume meiosis, are in the final stages of the first meiotic prophase and have already reached full size. If a spermatozoon penetrates this immature oocyte, it will fail to promote activation

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since the oocyte is not meiotically mature, and its chromosomes will undergo premature condensation22. GVBD may occur within minutes or require up to several hours after harvest; the length of time appears to depend on how far maturational events have progressed within the follicle before collection. More than 80% will succeed in passing through MI of maturation, ultimately to reach MII. The germinal vesicle, or nucleus, of the human oocyte is spherical and contains a large, refractile, exocentric nucleolus. Upon close examination, a second smaller nucleolus may be detected. The germinal vesicle is centrally located within the ooplasm of young PI oocytes and in those that exhibit developmental arrest. It migrates to a more cortical position in healthy oocytes before GVBD. The dissolution of the germinal vesicle marks the first practical microscopic indication that meiosis has resumed. As the oocyte matures, defenses against polyspermy are established in the form of cortical granule accumulation and alignment at the oocyte periphery. These granules are sparse and discontinuous in immature oocytes3. Under the microscope, the PI oocyte is characterized by its distinct germinal vesicle and refractile nucleolus. An irregular shape, darkened center and granular ooplasm are almost always displayed. Attached cumulus cells are usually compact and multilayered, but may be proliferative. Free follicular membrana granulosa cells within the immature follicle are usually small and appear in compact masses. PI oocytes with very mature characteristics of the cumulus (expanded appearance and very radiant corona) generally fail to undergo GVBD.

The sperm-penetrated human oocyte Fertilization process Human fertilization begins when a spermatozoon, with its haploid number of chromosomes, passes through oocyte cellular investments and makes contact with the protective zona pellucida that surrounds the oocyte. This contact induces an acrosomal reaction whereby the spermatozoon releases the contents of its acrosomal vesicle, including enzymes that aid the sperm in digesting its way through the zona to the oocyte plasma membrane. The equatorial segment of the sperm head attaches to the plasma membrane of the oocyte, and sperm incorporation occurs through a process similar to phagocytosis. Only acrosome-reacted sperm are believed to be capable of fusing with the oolemma of the oocyte. Spermatozoon-oolemma fusion is bypassed when performing intracytoplasmic sperm injection (ICSI) to assist the fertilization process. It has been reported that, under in vitro insemination conditions, spermatozoa transverse oocyte cellular investments by 3 h, and first appear in the oocyte cortex by 4 h. Of interest is that oocytes need to be incubated with spermatozoa for only 1 h to achieve fertilization outcomes similar to 16-h controls23. Fusion of gametes invokes a cascade of events that are initiated by the hydrolysis of phosphatidylinositol biphosphate in the oolemma24. Electrical changes occur on the oolemma, and intracellular calcium levels rise in the oocyte. Cortical granule exocytosis from the ooplasmic periphery causes a chemical alteration of the zona pellucida, which generally renders it impermeable to other sperm. Thus, the oocyte is said to become

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activated by its fusion with the spermatozoon. It completes its second meiotic division; 23 double-stranded chromosomes split at their centromeres, and chromatids separate to oocyte or second polar body. In this manner, a haploid number of chromosomes and a haploid amount of DNA are contributed by the oocyte. Activation does not necessarily require the stimulus of a spermatozoon. Oocytes can be activated through mechanical trauma, temperature shock, chemical stimulus or electrical signals. Oocytes are commonly activated during ICSI procedures by simply piercing the oolemma, or by aggressively disturbing the ooplasm. Within a few hours, male and female pronuclei are formed from the sperm and oocyte chromatin (Figure 1.7). The stage at which pronuclei are visible is termed the pronuclear stage, and the specimen is defined as a prezygote or ootid. Technically, the zygote has not yet formed (see Glossary). During pronuclear formation, the zygotic centrosome is assembled; centrosomal proteins and sperm aster microtubules gather around the sperm centriole. This assembly of the zygotic centrosome is a crucial step for subsequent pronuclear apposition and genomic union25. Pronuclei come in close contact, eventually lose their apposed pronuclear membranes and enter into syngamy (Figure 1.8). This final event of the fertilization process involves the reorganization and pairing of maternal and paternal chromosomes and formation of the zygote. Recall that the mixing of maternal and paternal gamete chromosomes during meiosis I results in a mathematical probability of more than eight million possible chromosome combinations (223) for each gamete. If each parent has this many combinations possible, a couple could produce more than 7×1013 offspring with different combinations of parental chromosomes. This astronomic number does not take into consideration the additional genetic variability generated by crossing-over events that occur during meiosis I. Without crossing-over, gene combinations on a given chromosome would remain coupled indefinitely. With crossing-over, the theoretical possibility of creating genetically different offspring after fertilization reaches 8023. This is why it is impossible, or nearly so, for any two individuals apart from monozygotic twins (or, in this age, clones) to be genetically identical. There is a brief period after pronuclei breakdown during which the zygote remains single-celled. In the human, the nearly 24-h long fertilization process is completed with the initiation of the first (mitotic) cleavage. Block to polyspermy One consequence of sperm-oolemma fusion is the exocytosis of cortical granules from the oocyte periphery. This release, occurring within minutes of fusion, is a key component of the oocyte’s strategy for preventing polyspermy. The dispersal of cortical granule contents into the perivitelline space is followed by a chemical alteration of the zona pellucida, an event often termed zona hardening or the zona reaction. Before fusion, the zona exhibits a porous appearance, and comprises a large number of ring-shaped structures called hoops, randomly superimposed in several layers; pore diameters decrease in size towards the inside of the zona. After fusion, the zona is observed to be more compact and its diameter decreases slightly; hoops are not distinguished and pores are obliterated by an amorphous material emerging from the inner zona26. The zona reaction may render the zona pellucida impenetrable by other sperm, or may cause

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secondary sperm to become entrapped in its altered matrix, unable to pass the highly condensed inner layer of the zona. A slow or incomplete cortical granule exocytosis and zona reaction may represent the most common causes of polyspermic fertilization. Premature or failed cortical granule discharge may be responsible for some instances of failed fertilization after standard insemination. It has been postulated that human oocytes do not possess a true block to polyspermy at the level of the oolemma27. This theory is supported by a study that retrospectively examined the polyspermy rate in over 3000 human oocytes subjected to subzonal insemination techniques (SUZI) when 1–20 sperm were placed under their zonae. The authors concluded that all sperm possessing fertilizing ability were indeed capable of fusing with the oocyte cell membrane, indicating the absence of a polyspermic block at this level28. However, in another experiment where zona-free human oocytes were exposed to high concentrations of sperm, it appeared that sperm were not able to penetrate oocytes indiscriminately at rates that would be expected29. At 30 min, anaverage of 1.3 sperm had penetrated the oocytes, and, at 60 min, 2.9 had been successful. The number of penetrating sperm peaked at 2 h, regardless of sperm concentration. In addition, sperm demonstrated a reduced ability to bind to membranes of previously fertilized oocytes, few or none binding to the membranes of 4-cell preembryos. These authors concluded that the oolemma does, in fact, play a role in preventing polyspermy and that a plasma membrane block may involve permanent changes to sperm binding/fusion ability. Based on these and other conflicting reports30, one can only conclude that, for the time being, the question of a membrane block in the human remains unresolved. Commonly, 5–10% of oocytes cultured in vitro are observed to incorporate more than one spermatozoon, as evidenced by the subsequent development of three or more pronuclei. The reported frequency in the literature ranges from as low as 1–2% after inseminating mature oocytes23,31 to greater than 30% after inseminating immature oocytes32. Some investigators have found a high correlation between triploidy and inseminating sperm concentration; as early as 1986, one such study reported a tripling of polyspermy with increasing sperm concentrations33. Others have reported that the incidence of abnormal fertilization is no higher when oocytes are exposed to large numbers of sperm15,34. Although it is tempting to try to correlate polyspermic fertilization to the unnaturally high numbers of spermatozoa used for standard insemination in vitro, it has been our observation that this incidence is better correlated to oocyte maturity and viability than to gross numbers of inseminating spermatozoa. Approximately 4–5% of mature oocytes exhibit three pronuclei after the injection of a single sperm during ICSI procedures (they are largely digynic), indicating a relatively high occurrence of second polar body suppression at meiosis II. Although one cannot dismiss the possibility that the ICSI procedure itself is instrumental in causing this, and although the possibility exists that a sperm possessing two nuclei was injected, digynic fertilization has been noted often enough after natural intercourse and in vitro insemination to suggest that it is not restricted to assisted fertilization techniques. Most late-term triploid fetuses and live-born triploid children have been shown to have developed from digynic preembryos35,36. Moreover, recent studies confirm that digyny is clearly the most common origin of triploidy in the human37. These studies indicate that oocyte factors are commonly accountable for triploid fertilization.

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Which oocyte factors might this include? Certainly, oocyte aging has been shown to be associated with an increased incidence of spindle defects; retention of chromosomes within the ooplasm after suppression of first or second polar body extrusion may represent one possible mechanism for digyny. Oocytes that are postmature (aged) have been shown to exhibit a centripetal migration of their cortical granules when analyzed under the electron microscope3,38. These would trigger a retarded zona reaction at best, which could result in multiple sperm fusion during fertilization. Conversely oocyte immaturity has been implicated in contributing to polyspermic fertilization, presumably due to delayed cortical granule release39. Poor in vitro culture conditions may be implicated in some cases of spindle damage if oocytes are allowed to chill for long periods40, or overheat. Additionally mature oocytes may develop from binucleate primary oocytes. In all probability polyspermy results from different mechanisms, or combinations of different mechanisms, in different oocytes. In some oocytes, immaturity or postmaturity may be implicated, or oocytes may be intrinsically abnormal. In others, minute cracks may be present in the zona pellucida after oocyte harvest procedures, allowing for multiple sperm entry. In others still, entry of two sperm may simply be a random event that occurs when two sperm simultaneously make their way through the zona and concurrently fuse with the oolemma; whether the odds for this happening increase in the presence of high numbers of motile spermatozoa remains to be elucidated. Male and female pronuclei Male and female pronuclei (Figure 1.9) are usually formed simultaneously; the male pronucleus forms near the site of sperm entry, and the female originates at the ooplasmic pole of the meiotic spindle41. These structures, although small and faint, may be visualized as early as 4 h after ICSI or 5–6 h after insemination. The male pronucleus may be somewhat larger in humans5, but the difference in some specimens is difficult, if not impossible, to discern under the light microscope. When one group of investigators attempted to distinguish pronuclear gender by using morphological criteria under the light microscope, they observed sperm tail remnants in only 3/342 pronucleate oocytes; furthermore, pronuclear diameter and position within the ooplasm failed to yield any informative distinctions between male and female pronuclei42. Early in their formation, pronuclei may be seen at a distance from each other; later, they migrate together towards the center of the cell. By 15 h after insemination, pronuclei are most often observed lying close to one another; they may present a ‘figure of 8’ appearance if viewed in an overlapping position. Although they appear to contact or fuse, transmission electron microscopy has demonstrated that they remain separated by a narrow strip of ooplasm that may contain mitochondria and elements of smooth endoplasmic reticulum, orbealtogether absent of organelles27,38,43. As male and female pronuclei become closely associated, adjacent areas of each appear to flatten out. During the same time, nucleoli move from random locations within each pronucleus to line up at the regions of juxtaposition. One to nine nucleoli can be observed in each structure, the smaller pronucleus often demonstrating a lower number. Pronuclei are surrounded by a dense aggregation of cellular organelles, which may appear granular or even darkened under the light microscope. The female pronucleus

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often dismantles its envelope and undergoes membrane breakdown slightly ahead of the male44. During the human pronuclear phase, DNA synthesis within male and female pronuclei begins synchronously at about 12 h after sperm-oocyte fusion. Errors of DNA synthesis may be responsible for developmental arrest at the pronuclear stage; it may be that pronuclear membranes require signalling of DNA replication before dismantling. In one elegant study, oocytes in the process of fertilization were monitored for up to 20 h by time-lapse video cinematography following ICSI45. Fertilization patterns in 50 oocytes followed a defined course of events, but varied markedly in timing between individual prezygotes. The investigators described a circular wave of granulation moving throughout the cytoplasm that lasted for approximately 20–53 min. Granulation occurred in cortical regions of the oocyte and moved in 2–10 full circular rotations, some clockwise, and some counter-clockwise. During this active phase, the sperm head decondensed. This was followed by extrusion of the second polar body and central development of the male pronucleus. After polar body extrusion (mean time from injection, 2.5 h), the granulation wave ceased in all oocytes. At the same time, or just after male pronucleus formation, the female pronucleus was seen to form near the site of second polar body extrusion, which was not always near the site of first polar body extrusion; it was gradually drawn towards the male pronucleus until the two abutted. Both pronuclei were then observed to increase in size and to contain moving nucleoli, some of which coalesced over time. During the period of pronuclear growth, cytoplasmic organelles were seen to migrate inwardly to the center of the oocyte, leaving a clear zone at the cortex. Measurements confirmed that the female pronucleus was indeed smaller than the male pronucleus in these specimens (22.4 µm vs. 24.1 µm, respectively), and possessed fewer nucleoli (4.2 vs. 7.0). It was discovered that subsequent preembryo quality, as judged by morphology and developmental rate, was correlated to sequential timing of events and duration of the cytoplasmic granulation wave, good preembryos showing uniform (though not necessarily more rapid) progression and longer granulation waves. It was interesting to note in this series that pronuclei could be identified as early as 3 h post-injection and that, by 5 h, over half of the oocytes possessed visible, small pronuclear structures. In this fascinating study, the existence of the cytoplasmic granulation wave proved to be a novel and unique finding. Without video cinematography, the embryologist must rely on the presence and number of pronuclei, assessed during one or two brief examinations, to determine whether or not normal fertilization is ongoing. Practical criteria for sperm penetration in living material include first, observation of two pronuclei at 10–18 h post-insemination, and second, visualization of two polar bodies in the perivitelline space. Assessment of these two parameters is rapid and simple. Unfortunately, the identification of two pronuclei cannot ensure a normal fertilization process and does not guarantee that one pronucleus is of paternal, and one is of maternal, origin. Evaluating second polar bodies is also potentially misleading because of first polar body fragmentation. Yet another serious drawback to using a single observation to assess pronuclear number lies in the fact that counts have been observed to change during the pronuclear period; most embryologists can relate instances of visualizing two pronuclei in an oocyte at an initial observation and one pronucleus (or three pronuclei) during a follow-up evaluation, or vice versa. While pronuclear and polar body determination is not ideal for the assessment of sperm penetration, it does provide the most useful and least time consuming means of

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clinical evaluation. Perhaps one day we will be evaluating all fertilized oocytes as described by Dianna Payne’s time-lapse video study45; until then, less informative methods will have to suffice. Chromosomes and fertilization Fusion between male and female gametes is not always successful, even under optimal conditions. When investigating causes of fertilization failure, gamete maturity and genetic health emerge as two important factors related to fertilizing potential. In one study carried out to examine fertilization failure in 293 oocytes inseminated in vitro, it was discovered that 30% of the oocytes were not fully mature at the time of sperm-oocyte interaction (chromosomes at MI or PI of maturation), and a full 59% were chromosomally abnormal46. Figures like these are often reported in the scientific literature, making it evident that a large proportion of human gametes are genetically incompetent to generate normal offspring. Gamete immaturity represents less of a problem now than it did when we began trying to optimize our IVF techniques many years ago. Certainly, ovarian stimulation regimes have been refined in the past 20 years, so that virtually all our patients produce healthy, mature oocytes. We have also been mildly successful in clinically applying in vitro maturation methods. Healthy babies have been generated from germinal vesicle-bearing immature oocytes collected from stimulated cycles20,47 and unstimulated cycles17,18. Unfortunately, implantation rates are not in the range expected from in vivo matured oocytes. Immature spermatozoa have generated considerable interest in recent years as well. Almost unthinkable two decades ago, the time has arrived when investigators are reporting the use of haploid round and elongated spermatids in the clinical treatment of azoospermia48,49, and healthy children have been conceived50,52. It now appears, based on experiments in the mouse, that injecting secondary spermatocytes will prove to be a future treatment modality; incredibly, normal offspring have been reported as being born after these cells completed meiosis II within oocytes, following injection and electroactivation. The extra set of chromosomes was extruded into the perivitelline space as an extra (male) polar body53. The fact that many gametes are genetically abnormal must account for much of the failed fertilization we observe in our programs. If we are to accept earnestly the many reports describing very high percentages of chromosome abnormalities in sperm, eggs and developing preembryos, it seems a wonder that we are managing so well to overpopulate the earth. It is well documented that chromosomal abnormalities among first-trimester spontaneous abortions occur at a rate of about 60%54. As well, it has been estimated that more than one-quarter of oocytes that fail fertilization55,56 and up to 10% of spermatozoa carry a chromosomal aberration57. A review of the literature led one investigator to conclude that at least 50% of conceptuses developing after natural conception are chromosomally abnormal58. Males with non-mosaic Klinefelter’s syndrome (47,XXY) are now capable of fathering children if even one or two mature or maturing sperm cells can be isolated from testicular tissue and used in assisted fertilization procedures. Whether the very presence of spermatozoa or spermatids in the testicular tissue indicates mosaicism in germ cell-

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lines must be investigated further. Several cases have been reported thus far showing normal karyotypes of preembryos generated from men without evidence of mosaicism in peripheral blood cells. In one report, after performing preimplantation genetic diagnosis on five preembryos from three Klinefelter’s individuals, all were found to be chromosomally normal59. In our own program, the first report of a Klinefelter’s birth was presented in 199860. A healthy, unaffected male child was delivered in this instance. Since then, seven other ongoing clinical pregnancies have been established using testicular spermatozoa from husbands with presumed non-mosaic Klinefelter’s syndrome. Three additional male children and seven female children have been delivered, inclusive of three twin sets. All of the children are healthy and perfectly normal in regard to their karyotypes. Through efforts such as these, we are continually learning more about the complex stages of reproduction. Perhaps no other branch of science is quite so interesting as the exploration of this fundamental life-generating process.

The cleaving human preembryo: 2-cell to 16-cell stage Cytokinesis Cytoplasmic division following nuclear replication and segregation is a universal characteristic of all cells. Cleavage of the human preembryo involves a series of mitotic divisions of the cytoplasm, every 12–18 h, without any discernible increase in its overall size (Figure 1.10). Failing to progress to the first cleavage after forming two pronuclei is relatively uncommon, occurring in less than or equal to 5% of normally fertilized oocytes61. As in most mammals, other than some rodents, the human sperm centrosome controls the first mitotic divisions after fertilization has taken place62. As the first cleavage mitosis reaches telophase, the cytoplasm of the zygote elongates and the surface contracts around the lesser circumference. This constriction continues until the zygote is divided into two blastomeres. The same process continues throughout all subsequent mitotic cell divisions21 (Figures 1.11 and 1.12). It has been estimated that mean blastomere volume is reduced by approximately 28.5% per division through the first three cleavages, and that some diversity normally exists between the volumes of sister blastomeres63. The 2–8-cell conceptus depends largely on the translation of stored maternal RNA for cleavage. Morphology The quality of 2–16-cell preembryos produced after IVF is variable. Many contain multiple cellular fragments or unequally sized blastomeres, or exhibit slow cell-doubling times. Fragments arise because blastomeres constantly change shape, making and breaking cell contacts during cleavage; in doing so they can leave behind cellular debris3. This constant, living motion is particularly apparent when specimens are viewed under time-lapse cinematography where continuous pulsing, formation of fragments and blebs, and cytoplasmic reorganization is noted (personal observation). Human and primate preembryos may be more disposed to these actions, since fragments are rarely seen in the conceptuses of other mammals. Because preembryos flushed from the uterine cavity after

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fertilization in vivo also exhibit fragments, it can be deduced that in vitro culture methods are not solely responsible for aberrant development. We certainly observe in our laboratories that preembryos exhibiting large numbers of anucleate fragments tend to implant less frequently. This may be the result of a reduction in the available cytoplasm for normal cell division that subsequently leads to reduced cell numbers in the blastocyst. Alternatively, numerous fragments may interfere with the process of compaction by making intimate cell-to-cell contact difficult. Cytoplasmic fragments often arise during the first cleavage. In studying the frequency of this observation at 24 h post-insemination, we found that when excessive numbers of fragments form so early, subsequent development of the preembryo is generally impaired. On the other hand, the development of small fragments after the first division usually has no detrimental effect on the cleaving conceptus. Extensive cytoplasmic fragmentation has been associated with impending preembryo death. After studying fragmented preembryos and comparing them to non-fragmented controls, Jurisicova and colleagues concluded that the high incidence of condensed chromatin, degraded DNA, cell corpses and apoptotic bodies commonly found in fragmented conceptuses almost certainly indicate a reduced potential for continued growth64. This contention is supported by the experience of most embryologists and physicians working in assisted reproduction programs, who observe lower implantation rates associated with irregular blastomeres and excessive fragmentation, although some preembryos with these qualities retain the capacity for implanting normally65. Shulman and co-workers reported that the implantation potential of transferred day-2 or -3 preembryos can be directly correlated to morphological parameters, and suggested that the number of preembryos replaced be balanced against their grade to reduce multiple gestation66. Rates of cleavage Many reports have linked normal cell-doubling times to preembryo viability, finding that slowly growing conceptuses demonstrate a markedly impaired capacity to implant after intrauterine transfer. The cell-doubling time in human preembryos between days 2 and 6 has been reported to be 31 h, with accelerated doubling noted after the first two divisions67. At first glance, these rates seem quite slow. During IVF treatment, one generally sees doubling in well under 24 h (2-cell stage by 24 h, 4-cell stage before 48 h, and 8-cell stage or more before 72 h), perhaps averaging every 18–20 h in healthy preembryos. However, it is not unreasonable to conclude that 31 h might represent the average doubling time if poor-quality and slowly growing and arrested preembryos are also calculated into the mean. In 1987, Claman and associates reported that 21/23 ongoing IVF pregnancies arose from transfers where at least one preembryo had reached the 4-cell stage by 40 h postinsemination68. Other reports have similarly concluded that preembryos with slow cleavage (fewer than four blastomeres at 42–44 h post-insemination) were less likely to produce a pregnancy69,70. McKiernan and Bavister demonstrated in the hamster that faster-cleaving preembryos not only lead to more morulae and blastocysts in culture, but that subsequent in vivo development of faster-growing conceptuses is associated with a higher incidence of viable fetuses71. These last authors suggest that the timely completion

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of the third cell cycle (8-cell stage) is a critical and favorable factor for predicting successful embryogenesis in the hamster. Regular cleavage to the 8-cell stage has also been noted as being a favorable observation in the human, but often proves to be an inexact tool for predicting implantation success when used as a single analytical parameter. Despite this, it has been proposed that developmental rate may be more important than morphology when weighing individual factors for human intrauterine transfer72. Some groups report that accelerated preembryonic growth combined with minimal fragmentation leads to increased pregnancy73. Yet other studies associate the occurrence of a timely first cleavage (before 25 h post-insemination) with enhanced pregnancy outcome74. Factors other than fragmentation and growth rate have also been associated with the implantation potential of human preembryos. These include zona pellucida thickness and/or variation in thickness, thin and variable being better75,77, adequate blastomere expansion76 and absence of multinucleation78. In addition, studies on follicular blood flow have demonstrated a high correlation between dissolved oxygen content in the follicle (greater than or equal to 3%) and subsequent normal development of the oocyte/preembryo79,81. On a practical basis, 2-cell conceptuses are observed any time after 20 h postinsemination, usually around 24 h, and may persist until 42 h post-insemination. Viable 4-cell preembryos are observed between 36 and 60 h post-insemination. Eight-cell stages are not generally seen until after 54 h, but usually before 72 h. In the human, seeing 3-, 5and 7-cell preembryos is not uncommon, particularly when the examination is carried out during mitotic cell division. This sometimes asynchronous division persists throughout cleavage of the early conceptus, and any number of blastomeres can be noted in a given observation. Interestingly, in some strains of mice and cows, male preembryos cleave faster than female ones82. There have been reports that this may be true for human preembryos as well83, although not all investigators have confirmed this finding84. In normally fertilized specimens, retarded growth (no doubling in 24 h) often indicates reduced viability, but accelerated cleavage (doubling in 12 h) may not necessarily reflect a healthier conceptus9. Occasionally, pregnancies are established with slowly growing preembryos, even those found to possess only 6–8 blastomeres at 96 h. In the Cornell program, intrauterine transfer is usually postponed for 1 day whenever preembryos are observed to possess fewer than five blastomeres on day 3. If any further cleavage occurs over the next 24 h, transfer is carried out; if no further cleavage occurs, transfer is cancelled. It has been surprising to note the number of pregnancies resulting from the transfer of 6- or 8-celled preembryos on day 4. A conceptus exhibiting three pronuclei may appear to cleave at an accelerated rate to the morula stage, at which time its development is usually arrested61. This is because many triploid zygotes split directly into three cells at the first cleavage because they possess a tripolar spindle; subsequent divisions reflect the higher overall cell number in the conceptus, not more rapid growth. Preembryo grading schemes (days 2–3 post-insemination) As a result of the reported correlations between morphology and pregnancy, embryologists generally use some sort of grading scheme to document the presumptive

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quality of transferred preembryos. Most of these schemes are related to the extent of observed cytoplasmic fragmentation and growth rate, but some include other factors such as zona pellucida thickness or blastomere size and regularity. Usually, a grade is assigned to the transfer based on the morphology of the highest-grade preembryo in the group, with additional fractions added or subtracted for appropriate growth or for the concurrent transfer of other exceptional conceptuses. Some groups will attempt to calculate an average score for the cohort of transferred preembryos, based on the assigned grades of each individual preembryo, but these systems tend to be less informative when wide disparity exists between conceptuses, distorting the meaningfulness of an averaged final figure. In one early scoring system, a morphological grade of 1–4 was given to each preembryo and then combined with a grade developed from direct comparison to ideal growth rate. Using this two-stage system, the scores proved to be of value in predicting clinical success85. Similarly, in 1987, Puissant and colleagues published the results of grading preembryos based on their number of anucleate fragments and rate of division86. It was found that those preembryos endowed with high grades contributed more often to pregnancy and multiple pregnancy. These authors recommended that, if grading scores are high in conjunction with optimal clinical parameters, fewer preembryos should be transferred to offset high multiple pregnancy rates. A three-grade scoring system was evaluated by Erenus and associates in 199187. Grade 1 preembryos represented those with equal-sized blastomeres and no fragmentation, grade 2 included preembryos with unequal-sized blastomeres and grade 3 included preembryos associated with cytoplasmic fragments. In cycles where the best preembryo transferred was grade 1, 22% achieved clinical pregnancy. This was compared to grades 2 and 3, where pregnancy rates were 13% and 0%, respectively. Additionally, pregnancy rates increased with the transfer of multiple grade 1 preembryos (40% with three grade 1 preembryos). In 1992, Steer and co-workers developed a cumulative grading system in an effort not only to predict pregnancy outcome, but also potentially to reduce high-order gestation in the Bourne Hall and Hallam programs88. With this system, the morphological grade of each preembryo (1–4; larger number associated with better morphology) was multiplied by the preembryo’s number of blastomeres. The sum of grades from all conceptuses transferred on day 2 after insemination represented the final score. It was retrospectively analyzed that pregnancy rates in women under age 36 years rose as the cumulative score increased to a maximum of 42. A continued increase above this number did not contribute further to establishing pregnancy, but did impact upon the multiple pregnancy rate. They estimated that, using this system prospectively, 78% of triplet and 100% of quadruplet pregnancies could have been predicted and avoided. Using this same system, Visser and Fourie reported pregnancy rates of only 4% associated with scores of 1–10, but greater than 35% with scores between 41 and 5089. They also found more biochemical pregnancies, but not clinical pregnancy losses, occurring with low-end scores. All triplet and quadruplet pregnancies were associated with scores above 40. Giorgetti and associates90 devised a system whereby preembryos were assigned one point for each of the following parameters: cleavage, no fragmentation, no irregular cells and four blastomeres on day 2. The idea for this system arose from the previous evaluation of 957 single preembryo transfers where no ongoing pregnancies were

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established from the transfer of uncleaved preembryos or after delayed fertilization (99 transfers), and where higher pregnancy rates were found to be associated with regular blastomeres and absence of fragmentation (858 transfers). Applying their four-point scoring system retrospectively, they discovered that both clinical pregnancy rates and delivery rates correlated significantly to a higher score after single preembryo transfer, and that each point corresponded to a 4% increase in pregnancy. Only female age (over 38 years) had as great an impact as the simple morphological assessment. Tasdemir and co-workers developed a system in which the degree of preembryo fragmentation and general morphology were assessed as being either good (A) or poor (B)91. They then examined the outcomes of transfers with AA, BB and AB double transfers, or AAA, BBB, AAB and ABB triple transfers. When only goodquality preembryos were transferred, the pregnancy rates in double (AA) and triple (AAA) transfers were 41% and 43%, respectively. When only poor-quality preembryos (BB and BBB) were transferred, rates were 11% and 23%. AB transfers resulted in a 37% pregnancy rate, and AAB/ABB transfers resulted in a 40% incidence. The authors concluded that if at least one good-quality preembryo was available for transfer, then double, rather than triple, transfer should be carried out; higher numbers should be considered only in cases of poor preembryo quality. After applying this policy for 1 year in patients under the age of 37, they compared their results to previous data generated from triple transfers. Although this prospective trial demonstrated a slightly lower pregnancy rate in the study group as compared to the control group, the authors believe that the lower incidence of triplet gestations provides a practical compromise between high pregnancy rates and high-order gestation92. At Cornell, a system is used first to grade the morphology of cleaving preembryos and second, to classify transfers according to the highest score in the cohort of conceptuses being replaced: Grade 1 preembryo with blastomeres of equal size; no cytoplasmic fragmentation; Grade 2 preembryo with blastomeres of equal size; minor cytoplasmic fragmentation covering less than or equal to 15% of the preembryo surface; Grade 3 preembryo with blastomeres of distinctly unequal size; variable fragmentation; Grade 4 preembryo with blastomeres of equal or unequal size; moderate to significant cytoplasmic fragmentation covering greater than or equal to 20% of the preembryo surface; Grade 5 preembryo with few blastomeres of any size; severe fragmentation covering greater than or equal to 50% of the preembryo surface. We have observed that transfers with at least one grade 1 or grade 2 preembryo possess a greater potential for establishing pregnancy93. When data are normalized for the number of preembryos transferred, this trend still exists for all groups except single preembryo transfer, where the number of replacements is too low for comparison and the

An atlas of human blastocysts 18

group is highly represented by patients with poor ovarian response. Clearly, transferring three or four preembryos of good quality produces the best clinical pregnancy rates, albeit with a concurrent increase in multiple implantations. Although a higher score (lower number) is favorable, pregnancy is quite possible even in cycles with grade 4 or 5 morphology demonstrating unequal-sized blastomeres and moderate to severe cytoplasmic fragmentation. Of interest is that scores are remarkably repetitive for the same patient in succeeding cycles. In the Cornell program, two or three day-3 (6-cell to 10-cell) preembryos are replaced in women under the age of 34; three or four are recommended for women between the ages of 34 and 39; five are often replaced in women over the age of 40 when and if they are available; more may be considered in special circumstances over the age of 43 years. This strategy is based on the obstetrical outcomes of more than 7000 IVF deliveries. While the multiple pregnancy rate is high when more than two are replaced, particularly in young women, the vast majority are twin gestations, suitable to most infertile couples. An attractive alternative is to replace one less preembryo than described above in cycles producing adequate numbers of preembryos with grade 1 or 2 morphology. Doing so may result in continued acceptable pregnancy rates while reducing the occurrence of multiples (see Chapter 6). It has been reported that the concept of older women establishing multiple pregnancy at lower rates than younger women is a fallacy94. In this report, the authors suggest limiting the number of preembryos for transfer to three, regardless of age. Our own data do not support reducing transfer numbers in this particular population. After examining replacement outcomes during 6 years for 1876 cycles involving women over age 40, we find a significantly higher clinical pregnancy rate per transfer when four or more preembryos are replaced as compared to three (43% vs. 28%, respectively; p<0.0001), as well as a significant difference in the incidence of multiple pregnancy per transfer (11% vs. 6%). One could argue that, since the policy exists to replace more than three preembryos in older women whenever possible, those receiving three represent women with poor ovarian response or limited oocyte reserve; this is quite true and should be overlooked when weighing the comparisons. Nevertheless, transferring more than three preembryos to older patients does not appear to expose these women to excessively or unacceptably high rates of multiple pregnancy, and does not lead to the multiple pregnancy rates seen after transfer of three or more preembryos to younger women (three replaced under age 40, 61% clinical pregnancy and 28% multiple pregnancy; more than three replaced under age 40, 58% clinical pregnancy and 27% multiple pregnancy). The selection of viable preembryos for intrauterine transfer is considered an important factor in establishing pregnancy; it is assumed that overall morphology and cleavage rate will, at least to some extent, reflect a preembryo’s potential for continued growth and implantation (see Chapter 6). In deciding how many preembryos to transfer to a given individual on day 2 or 3, it seems most reasonable to weigh the risks of multiple pregnancy against age, previous history, preembryo morphology and development, desires of the couple, and the delivery rates of a given clinic.

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Totipotency As discussed in later chapters, individual blastomeres remain distinct, and are totipotent (capable of developing independently to form a new organism) until about the 8-cell stage, when changes occur in the structure and properties of their plasma membranes and cytoplasm. Cells at this point become less distinct as they adhere more tightly to one another during the process of compaction. After compaction, they begin to commit themselves to becoming either inner cell mass or trophectoderm, and thus, lose this totipotency.

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An atlas of human blastocysts 20 15. Veeck LL. Atlas of the Human Oocyte and Early Conceptus, 2nd edn. Baltimore: Williams & Wilkins, 1991:3, 13, 27 (granulosa), 13 (oocyte classification), 27 (disparity cumulus and nucleus, 218 (spermatozoa) 16. Coetzee K, Windt ML. Fertilization and pregnancy using metaphase I oocytes in an intracytoplasmic sperm injection program.J Assist Reprod Genet 1996; 13:768–71 17. Barnes FL, Crombie A, Gardner DK, et al. Blastocyst development and birth after in-vitro maturation of human primary oocytes, intracytoplasmic sperm injection and assisted hatching. Hum Reprod 1995; 10:3243–7 18. Cha KY, Koo JJ, Ko JJ, Choi DH, Han SY, Yoon TK. Pregnancy after in vitro fertilization of human follicular oocytes collected from nonstimulated cycles, their culture in vitro and their transfer in a donor oocyte program. Fertil Steril 1991; 55:109–13 19. Liu J, Katz E, Garcia JE, Compton G, Baramki TA. Successful in vitro maturation of human oocytes not exposed to human chorionic gonadotropin during ovulation induction, resulting in pregnancy. Fertil Steril 1997; 67:566–8 20. Veeck LL, Wortham JW Jr, Witmyer J, et al. Maturation and fertilization of morphologically immature human oocytes in a program of in vitro fertilization. Fertil Steril 1983; 39:594–602 21. Austin CR. The Mammalian Egg. Oxford: Blackwell Scientific Publications, 1961:75, 78 22. Plachot M, Crozet N. Fertilization abnormalities in human in-vitro fertilization. Hum Reprod 1992;7(Suppl 1):89–94 23. Gianaroli L, Cristina Magli M, Ferraretti AP, et al. Reducing the time of sperm-oocyte interaction in human in-vitro fertilization improves the implantation rate. Hum Reprod 1996; 11:166–71 24. Alberts B, Bray D, Lewis J, Raff M, Roberts K, Watson JD, eds. Germ cells and fertilization. In Molecular Biology of the Cell, 2nd edn. New York: Garland Publishing, 1989:877 25. Sutovsky P, Hewitson L, Simerly C, Schatten G. Molecular medical approaches for alleviating infertility and under standing assisted reproductive technologies. Proc Assoc Am Physicians 1996; 108:432–43 26. Nikas G, Paraschos T, Psychoyos A, Handyside AH. The zona reaction in human oocytes as seen with scanning electron microscopy. Hum Reprod 1994; 9:2135–8 27. Soupart P, Strong PA. Ultrastructural observations on human oocytes fertilized in vitro. Fertil Steril 1974; 25:11–44 28. Wolf JP, Ducot B, Aymar C, et al. Absence of block to polyspermy at the human oolemma. Fertil Steril 1997; 67:1095–102 29. Sengoku K, Tamate K, Horikawa M, Takaoka Y, Ishikawa M, Dukelow WR. Plasma membrane block to polyspermy in human oocytes and preimplantation embryos. J Reprod Fertil 1995; 105:85–90 30. Van Blerkom J, Davis PW, Merriam J. The developmental ability of human oocytes penetrated at the germinal vesicle stage after insemination in vitro. Hum Reprod 1994; 9:697–708 31. Van Blerkom J, Henry G, Porreco R. Preimplantation human embryonic development from polypronuclear eggs after in vitro fertilization. Fertil Steril 1984; 41:686–96 32. van der Ven HH, AI-Hasani S, Diedrich K, Hamerich U, Lehmann F, Krebs D. Polyspermy in in vitro fertilization of human oocytes: frequency and possible causes. Ann NY Acad Sci 1985; 442:88–95 33. Englert Y, Puissant F, Camus M, Degueldre M, Leroy F. Factors leading to tripronucleate eggs during human in-vitro fertilization. Hum Reprod 1986; 1:117–19 34. Diamond MP, Rogers BJ, Webster BW, Vaughn WK, Wentz AC. Polyspermy: effect of varying stimulation protocols and inseminating sperm concentrations. Fertil Steril 1985; 43:777–80 35. Dietzsch E, Ramsay M, Christianson AL, Henderson BD, de Ravel TJ. Maternal origin of extra haploid set of chromosomes in third trimester triploid fetuses. Am J Med Genet 1995; 58:360–4 36. Miny P, Koppers B, Dworniczak B, et al. Parental origin of the extra haploid chromosome set in triploidies diagnosed prenatally. Am J Med Genet 1995; 57:102–6

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37. McFadden DE, Pantzar JT. Placental pathology of triploidy. Hum Pathol 1996; 27:1018–20 38. Sathananthan AH. Ultrastructural morphology of fertilization and early cleavage in the human. In Trounson AO, Wood C, eds. In Vitro Fertilization and Embryo Transfer. London: Churchill Livingstone, 1984:110 (pronuclei), 131 (cortical granules) 39. Sathananthan AH, Trounson AO. Ultrastructure of cortical granule release and zona interaction in monospermic and polyspermic human ova fertilized in vitro. Gamete Res 1982; 6:225 40. Almeida PA, Bolton VN. The effect of temperature fluctuations on the cytoskeletal organisation and chromosomal constitution of the human oocyte. Zygote 1995; 3:357–65 41. Wright G, Wiker S, Elsner C, et al. Observations on the morphology of pronuclei and nucleoli in human zygotes and implications for cryopreservation. Hum Reprod 1990; 5:109–15 42. Wiker S, Malter H, Wright G, Cohen J. Recognition of paternal pronuclei in human zygotes. J In Vitro Fertil Embryo Transf1990; 7:33–7 43. Zamboni L. In Fine Morphology of Mammalian Fertilization, 1st edn. New York: Medical Department, Harper & Row, 1971 44. Sathananthan AH, Trounson AO. The human pronuclear ovum: fine structure of monospermic and polyspsermic fertilization in vitro. Gamete Res 1985; 12:385 45. Payne D, Flaherty SP, Barry MF, Matthews CD. Preliminary observations on polar body extrusion and pronuclear formation in human oocytes using time-lapse video cinematography. Hum Reprod 1997; 12:532–41 46. Almeida PA, Bolton VN. Immaturity and chromosomal abnormalities in oocytes that fail to develop pronuclei following insemination in vitro. Hum Reprod 1993; 8:229–32 47. Nagy ZP, Cecile J, Liu J, Loccufier A, Devroey P, Van Steirteghem A. Pregnancy and birth after intracytoplasmic sperm injection of in vitro matured germinal-vesicle stage oocytes: case report. Fertil Steril 1996; 65:1047–50 48. Chen SU, Ho HN, Chen HF, Tsai TC, Lee TY, Yang YS. Fertilization and embryo cleavage after intracytoplasmic spermatid injection in an obstructive azoospermic patient with defective spermiogenesis. Fertil Steril 1996; 66:157–60 49. Vanderzwalmen P, Lejeune B, Nijs M, Segal-Bertin G, Vandamme B, Schoysman R. Fertilization of an oocyte microinseminated with a spermatid in an in-vitro fertilization programme. Hum Reprod 1995; 10:502–3 50. Fishel S, Green S, Bishop M, et al. Pregnancy after intra-cytoplasmic injection of spermatid. Lancet 1995; 345:1641–2 51. Tesarik J, Mendoza C, Testart J. Viable embryos from injection of round spermatids into oocytes. N Engl J Med 1995; 333:525 52. Antinori S, Versaci C, Dani G, Antinori M, Pozza D, Selman HA. Fertilization with human testicular spermatids: four successful pregnancies. Hum Reprod 1997; 12:286–91 53. Kimura Y, Yanagimachi R. Development of normal mice from oocytes injected with secondary spermatocyte nuclei. Biol Reprod 1995; 53:855–62 54. Boue J, Bou A, Lazar P. Retrospective and prospective epidemiological studies of 1500 karyotyped spontaneous human abortions. Teratology 1975; 12:11–26 55. Plachot M. Cytogenetic analysis of oocytes and embryos. Ann Acad Med Singapore 1992; 21:538–44 56. Plachot M. The human oocyte. Genetic aspects. Ann Genet 1997; 40:115–20 57. Martin RH, Balkan W, Burns K, Rademaker AW, Lin CC, Rudd NL. The chromosome constitution of 1000 human spermatozoa. Hum Genet 1983; 63:305–9 58. Schulman JD, Dorfmann A, Evans MI. Genetic aspects of in vitro fertilization. Ann NYAcad Sci 1985; 442:466–75 59. Staessen C, Coonen E, Van Assche E, et al. Preimplantation diagnosis for X and Y normality in embryos from three Klinefelter patients. Hum Reprod 1996; 11:1650–3 60. Palermo GD, Schlegel PN, Sills ES, et al. Births after intra-cytoplasmic injection of sperm obtained by testicular extraction from men with nonmosaic Klinefelter’s syndrome. N Engl J Med 1998; 338:588–90

An atlas of human blastocysts 22 61. Van Blerkom J. Developmental failure in human reproduction associated with preovulatory oogenesis and preimplantation embryogenesis. In Van Blerkom J, Motta PM, eds. Ultrastructure of Human Gametogenesis and Early Embryogenesis; Electron Microscopy in Biology and Medicine 5. Boston: Kluwer Academic Publishers, 1989:125 62. Palermo G, Munne S, Cohen J. The human zygote inherits its mitotic potential from the male gamete. Hum Reprod 1994; 9:1220–5 63. Goyanes VJ, Ron-Corzo A, Costas E, Maneiro E. Morphometric categorization of the human oocyte and early conceptus. Hum Reprod 1990; 5:613–18 64. Jurisicova A, Varmuza S, Casper RF. Programmed cell death and human embryo fragmentation. Mol Hum Reprod 1996; 2:93–8 65. Grillo JM, Gamerre M, Lacroix O, Noizet A, Vitry G. Influence of the morphological aspect of embryos obtained by in vitro fertilization on their implantation rate. J In Vitro Fertil Embryo Transf 1991; 8:317–21 66. Shulman A, Ben-Nun I, Ghetler Y, Kaneti H, Shilon M, Beyth Y. Relationship between embryo morphology and implantation rate after in vitro fertilization treatment in conception cycles. Fertil Steril 1993; 60:123–6 67. Herbert M, Wolstenholme J, Murdoch AP, Butler TJ. Mitotic activity during preimplantation development of human embryos. J Reprod Fertil 1995; 103:209–14 68. Claman P, Armant DR, Seibel MM, Wang TA, Oskowitz SP, Taymor ML. The impact of embryo quality and quantity on implantation and the establishment of viable pregnancies. J In Vitro Fertil Embryo Transf 1987; 4:218–22 69. Lewin A, Schenker JG, Safran A, et al. Embryo growth rate in vitro as an indicator of embryo quality in IVF cycles. J Assist Reprod Genet 1994; 11:500–3 70. Zhu J, Meniru Gl, Craft IL. Embryo developmental stage at transfer influences outcome of treatment with intracytoplasmic sperm injection. J Assist Reprod Genet 1997; 14:245–9 71. McKiernan SH, Bavister BD. Timing of development is a critical parameter for predicting successful embryogenesis. Hum Reprod 1994; 9:2123–9 72. Ziebe S, Petersen K, Lindenberg S, Andersen AG, Gabrielsen A, Andersen AN. Embryo morphology or cleavage stage: how to select the best embryos for transfer after in-vitro fertilization. Hum Reprod 1997; 12:1545–9 73. Wiemer KE, Dale B, Hu Y, Steuerwald N, Maxson WS, Hoffman Dl. Blastocyst development in co-culture: development and morphological aspects. Hum Reprod 1995; 10:3226–32 74. Shoukir Y, Campana A, Farley T, Sakkas D. Early cleavage of in-vitro fertilized human embryos to the 2-cell stage: a novel indicator of embryo quality and viability. Hum Reprod 1997; 12:1531–6 75. Cohen J, Inge KL, Suzman M, Wiker SR, Wright G. Videocinematography of fresh and cryopreserved embryos: a retrospective analysis of embryonic morphology and implantation. Fertil Steril 1989; 51:820–7 76. Morgan K, Wiemer K, Steuerwald N, Hoffman D, Maxson W, Godke R. Use of videocinematography to assess morphological qualities of conventionally cultured and cocultured embryos. Hum Reprod 1995; 10:2371–6 77. Garside WT, Loret de Mola JR, Bucci JA, Tureck RW, Heyner S. Sequential analysis of zona thickness during in vitro culture of human zygotes: correlation with embryo quality, age, and implantation. Mol Reprod Dev 1997; 47:99–104 78. Kligman I, Benadiva C, Alikani M, Munne S. The presence of multinucleated blastomeres in human embryos is correlated with chromosomal abnormalities. Hum Reprod 1996; 11:1492–8 79. Nargund G, Bourne T, Doyle P, et al. Associations between ultrasound indices of follicular blood flow, oocyte recovery and preimplantation embryo quality. Hum Reprod 1996; 11:109–13 80. Chui DK, Pugh ND, Walker SM, Gregory L, Shaw RW. Follicular vascularity—the predictive value of transvaginal power Doppler ultrasonography in an in-vitro fertilization programme: a preliminary study. Hum Reprod 1997; 12:191–6

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81. Van Blerkom J. Can the developmental competence of early human embryos be predicted effectively in the clinical IVF laboratory? Hum Reprod 1997; 12:1610–14 82. Leese HJ, Edwards RG. The potential for preimplantation diagnosis by non-invasive methods. In Edwards RG, ed. Preconception and Preimplantation Diagnosis. Cambridge: Cambridge University Press, 1993:299 83. Ray PF, Conaghan J, Winston RM, Handyside AH. Increased number of cells and metabolic activity in male human preimplantation embryos following in vitro fertilization. J Reprod Fertil 1995; 104:165–71 84. Ng E, Claman P, Leveille MC, et al. Sex ratio of babies is unchanged after transfer of fastversus slow-cleaving embryos. J Assist Reprod Genet 1995; 12:566–8 85. Cummins JM, Breen TM, Harrison KL, Shaw JM, Wilson LM, Hennessey JF. A formula for scoring human embryo growth rates in in vitro fertilization: its value in predicting pregnancy and in comparison with visual estimates of embryo quality. J In Vitro Fertil Embryo Transf 1986; 3:284–95 86. Puissant F, Van Rysselberge M, Barlow P, Deweze J, Leroy F. Embryo scoring as a prognostic tool in IVF treatment. Hum Reprod 1987; 2:705–8 87. Erenus M, Zouves C, Rajamahendran P, Leung S, Fluker M, Gomel V. The effect of embryo quality on subsequent pregnancy rates after in vitro fertilization. Fertil Steril 1991; 56:707–10 88. Steer CV, Mills CL, Tan SL, Campbell S, Edwards RG. The cumulative embryo score: a predictive embryo scoring technique to select the optimal number of embryos to transfer in an in-vitro fertilization and embryo transfer programme. Hum Reprod 1992; 7:117–19 89. Visser DS, Fourie FR. The applicability of the cumulative embryo score system for embryo selection and quality control in an in-vitro fertilization/embryo transfer programme. Hum Reprod 1993; 8:1719–22 90. Giorgetti C, Terriou P, Auquier P, et al. Embryo score to predict implantation after in-vitro fertilization: based on 957 single embryo transfers. Hum Reprod 1995; 10:2427–31 91. Tasdemir M, Tasdemir I, Kodama H, Fukuda J, Tanaka T. Two instead of three embryo transfer in in-vitro fertilization. Hum Reprod 1995; 10:2155–8 92. Kodama H, Fukuda J, Karube H, et al. Prospective evaluation of simple morphological criteria for embryo selection in double embryo transfer cycles. Hum Reprod 1995; 10:2999–3003 93. Veeck LL. An Atlas of Human Gametes and Conceptuses: an lllustrated Reference for Assisted Reproductive Technology. Carnforth, UK: Parthenon Publishing Group, 1999 94. Senoz S, Ben-Chetrit A, Casper RF. An IVF fallacy: multiple pregnancy risk is lower for older women. J Assist Reprod Genet 1997; 14:192–8

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Figure 1.1 Mature oocytes at metaphase II of maturation. (a) A mature oocyte in its natural state near the time of ovulation. At ×100, the

Overview of early human preimplantation Overview of early human preimplantation

oocyte is surrounded by both an expanded corona radiata which is many layers thick, and a dense outer cumulus oophorus. Under the cover of obscuring cells, the oocyte itself is somewhat difficult to visualize; nonetheless, a first polar body was confirmed after enzymatic removal of cells; (b) MII oocyte exhibiting spherical shape and first polar body at a 12-o’clock position; (c) MII oocyte possessing minor perivitelline debris; first polar body at 8-o’clock position; (d) MII oocyte; first polar body at 6o’clock position

Figure 1.2 Fluorescence photomicrograph of a normal meiotic spindle of an oocyte with chromosomes at metaphase II of maturation; stained with α-tubulinFITC (green); small insert to bottom right shows DNA lined up on the

25

An atlas of human blastocysts 26

equatorial plane, stained with DAPI (blue)

Figure 1.3 Oocytes at metaphase I of maturation. (a) Spherical, late metaphase I oocyte with perivitelline debris; zona pellucida is distinguished by two distinct layers; (b) spherical, late metaphase I oocyte with zona pellucida that demonstrates variable thickness; (c) spherical, late metaphase I oocyte; large perivitelline space; (d) earlier metaphase I oocyte as compared to (a)–(c); shape is irregular; zona pellucida demonstrates bilayering defect at right

Overview of early human preimplantation Overview of early human preimplantation

Figure 1.4 Fluorescence photomicrograph of a disorganized meiotic spindle of an aged oocyte; stained with α-tubulin-FITC (green). DNA, stained with DAPI (blue), is scattered and chaotic

Figure 1.5 Oocytes exhibiting a triploid condition after in vitro aging. (a) Oocyte after ICSI displaying three pronuclei; the fact that one pronucleus is large and two are smaller suggests that retention of polar body

27

An atlas of human blastocysts 28

chromosomes may be the cause of abnormal fertilization (digyny). In fact, polar bodies cannot be identified clearly; (b) inseminated oocyte with three pronuclei; the fact that two pronuclei are large and one is smaller suggests that penetration by more than one spermatozoon or penetration by a binucleate spermatozoon may be the cause of abnormal fertilization (diandry). Nucleoli in the bottom pronucleus and the one to the left appear abnormal in regard to number

Figure 1.6 Oocytes at prophase I of maturation. (a) Typical germinalvesicle (GV)-bearing oocyte showing coarse granularity, exocentric GV, and extremely thick zona pellucida; (b)

Overview of early human preimplantation Overview of early human preimplantation

immature oocyte with irregular shape and a clear peripheral zone; germinal vesicle is located at 3-o’clock; (c) immature oocyte with exocentric germinal vesicle; (d) oocyte with centrally located germinal vesicle displaying extreme granularity and exaggerated clear peripheral zone

29

An atlas of human blastocysts 30

Figure 1.7 Sperm-penetrated oocytes displaying two pronuclei. (a) Pronuclei of approximately equal size; nucleoli beginning to align at adjacent areas within pronuclei; (b) smallest

Overview of early human preimplantation Overview of early human preimplantation

pronucleus nearest polar bodies; female pro-nucleus has aligning nucleoli while male pronucleus retains scattered nucleoli; zona pellucida thick; (c) clear cortical zone and coarse granularity of cytoplasm; pronuclei of approximately equal size and shape; (d) clear cortical zone; female pronucleus nearest two polar bodies, one of which has fragmented; (e) female pronucleus nearest polar bodies and contains a larger number of smaller nucleoli; male pronucleus contains three nucleoli

Figure 1.8 Fertilized oocytes in syngamy. (a) At 20 hours post-

31

An atlas of human blastocysts 32

insemination, pronuclei have disappeared entirely; (b) at 20 hours post-insemination, a faint remnant of one central pronucleus is still seen

Figure 1.9 Male and female pronuclei; high-power magnification. (a) Fertilizing oocyte displaying two pronuclei and two polar bodies; nucleoli are aligned at adjacent pronuclear interfaces; (b) pronuclear oocyte; non-aligned nucleoli of differing sizes; (c) pronuclear oocyte; female pronucleus nearest polar bodies and differing nucleoli sizes; (d)

Overview of early human preimplantation Overview of early human preimplantation

pronuclear oocyte with pronuclei of distinctly different sizes; clear cortical zone

Figure 1.10 Preeembryos after the first cleavage (2-cell stage of development).

33

An atlas of human blastocysts 34

(a) First cleavage in progress; (b)–(e) two-cell stages

Figure 1.11 Preembryos after the second cleavage (4-cell stage of development)

Overview of early human preimplantation Overview of early human preimplantation

Figure 1.12 Preembryos at the 6–8cell stages of development. (a) Six-cell stage conceptus displaying equal-sized blastomeres and healthy appearance; (b) 8-cell stage; very minor fragmentation along periphery; (c) 8cell stage; thin zona pellucida; no fragmentation; (d) two preembryos before transfer, 7-cell stage to left and 8-cell to right; (e) and (f) preembryos with 12 blastomeres

35

2 Metabolic requirements during preimplantation development and the formulation of culture media David K.Gardner and Michelle Lane

Introduction Conventional culture conditions for the preimplantation mammalian embryo induce a considerable amount of cellular stress within the blastomeres of the embryo. Manifestations of such stress are retarded rates of cleavage, cleavage arrest, cytoplasmic blebbing, abnormal genome activation and gene transcription, and abnormal patterns of energy metabolism. All of these manifestations of culture-induced stress culminate in a loss of viability. It is therefore paramount that culture media minimize such stress within the embryo and facilitate normal cell function, and hence maintain embryo viability1–3. One of the cellular processes affected by suboptimal culture conditions, energy metabolism, has been shown to be a useful marker of embryo normalcy in vitro. Furthermore, energy metabolism may also be used to identify the most viable embryo from within a given cohort for transfer. It is therefore the aim of this chapter not only to review the nutrient requirements and energy metabolism of the mammalian preimplantation embryo, but also to put energy metabolism into a meaningful context with regard to embryo development, culture and viability assessment.

Nutrient requirements of the embryo The preimplantation period of human embryo development is highly dynamic and takes place in a changing environment in vivo4. Table 2.1 highlights some of the major differences in the embryo before and after compaction. With regard to carbohydrate utilization, the zygote and cleavage-stage embryo consume pyruvate

Table 2.1 Differences in embryo physiology preand post-compaction Pre-compaction

Post-compaction

Low biosynthetic activity

high biosynthetic activity

Low QO2

high QO2

Pyruvate preferred nutrient

glucose preferred nutrient

An atlas of human blastocysts 38

Non-essential amino acids

non-essential+essential amino acids

Maternal genome

embryonic genome

Individual cells

transporting epithelium

One cell type

two distinct cell types: ICM and trophectoderm

QO2, oxygen consumption/mg protein; ICM, inner cell mass

and lactate preferentially5,6, while post-compaction the embryo switches to a more glucose-based metabolism5–7. The reasons and mechanisms underlying this biphasic metabolism are described in detail below. Along with changes in carbohydrate utilization, the preimplantation embryo also undergoes changes in its requirements for amino acids8,9. Prior to compaction, embryo development is highest in the presence of alanine, aspartate, asparagine, glutamate, glutamine, glycine, proline, serine and taurine10, while after compaction the embryo’s requirements for amino acids increase, and development and differentiation are supported by a wider array of amino acids8. The roles of amino acids in embryo development are listed in Table 2.2. It is proposed that amino acids are among the most important regulators of gamete and embryo function, and the use of any culture medium lacking amino acids will confer significant trauma to the embryo3.

Table 2.2 Functions of amino acids during preimplantation mammalian embryo development Function

Reference

Biosynthetic precursors

11

Sources of energy

12

Regulators of energy metabolism

13

Osmolytes

14

Buffers of intracellular pH

15

Antioxidants

16

Chelators

17

Energy metabolism during the preimplantation period The mammalian preimplantation embryo undergoes significant changes in the way in which it generates its cellular energy as it develops from the zygote to the blastocyst stage1,18–20. During development, the relative activities of the two major energygenerating pathways, glycolysis and the tricarboxylic acid (TCA) cycle, change dramatically, to the extent that the zygote and blastocyst are metabolically analogous to two totally different somatic cell types. The zygote, and the oocyte from which it was derived, have a relatively low oxygen consumption21–23, and depend on low levels of

Metabolic requirements during preimplantation development

39

oxidation of pyruvate and/or lactate20,24. The zygote cannot use glucose as the sole energy source25. In contrast, the blastocyst exhibits a large capacity to utilize glucose both oxidatively26 and through aerobic glycolysis27 (aerobic glycolysis is here defined as the conversion of glucose to lactate even in the presence of adequate levels of oxygen for oxidative metabolism). As such, the zygote can be likened to a quiescent tissue such as bone, while the blastocyst to an invasive tumor. The significance of these similarities to other tissues is reflected in the physiology of the embryos at these given times. Whereas the oocyte remains relatively dormant in the ovary for a significant percentage of the female’s life, the blastocyst not only has to proliferate and differentiate, but also has to maintain the blastocoel and subsequently invade the endometrium. Of significance is that the levels of nutrients available to the human embryo as it develops mirror the changing requirements of the embryo4. At the time when the cleavage-stage embryo is present in the oviduct, the fluid is characterized by relatively high levels of pyruvate and lactate and low levels of glucose. In contrast, the fluid within the uterus is characterized by relatively high levels of glucose and low levels of pyruvate and lactate (Table 2.3). Furthermore, the cumulus cells surrounding the oocyte, zygote and early cleavage stage embryo readily consume glucose and convert it to both pyruvate and lactate4,28. Subsequently the human embryo will be exposed to a declining pyruvate and lactate gradient, but an increasing glucose gradient, as it develops.

Understanding the change in carbohydrate preference and utilization The change in carbohydrate utilization can be explained in terms of the changing physiology of the embryo as it develops. The oocyte, and hence the zygote, exhibit relatively low levels of biosynthesis prior to embryonic genome activation and expression. As a direct result of this there is a high adenosine triphosphate/adenosine diphosphate (ATP/ADP) ratio within the embryo, which in turn will allosterically inhibit the flux of glucose through the glycolytic pathway19,29,30. As the embryo becomes increasingly transcriptionally active31, protein synthesis increases32, and as the blastocoel is formed

Table 2.3 Concentration of carbohydrates in the human oviduct and uterus. Data from reference 4 Pyruvate (mmol/l)

Lactate* (mmol/l)

Glucose (mmol/l)

Oviduct (mid-cycle)

0.32

10.50

0.50

Uterus

0.10

5.87

3.15

*Lactate measured as the biologically active L-isoform

through the action of the basolateral ATPases33,34, there will be an increasing demand for energy (ATP). Consequently, the ATP/ADP ratio in the later-stage embryo will fall35,36, and an increased glycolytic flux will become possible. Indeed, glucose metabolism by the

An atlas of human blastocysts 40

mammalian embryo increases with development, with the highest rates of utilization occurring at the blastocyst stage12,37–40. Alternative explanations proposed for the observed switch in carbohydrate utilization are the lack of glucose carriers on the plasma membrane of the early embryo and/or the synthesis of sufficient enzymes for glucose metabolism by the later stages41. However, it is unlikely that these are the underlying mechanisms involved in the switch, as a carrier for glucose appears to present at all stages of development42–45, and there appears to be more than sufficient total enzyme activity present to accommodate glucose metabolism through glycolysis46,47. What is probably more important than total enzyme activity is the appearance of new isoforms of enzymes as development proceeds. The significance of enzyme isoforms and their relative changes in abundance in terms of embryo physiology/metabolism has not been fully determined, although it is evident that the isoforms of key enzymes do change during the pre- and peri-implantation period1,48,49. Different isoforms vary in their response to specific regulators of enzyme function, and could therefore help to explain the observed metabolic responses of different stages of development to their environment in culture3.

Is glucose toxic to the embryo? In simple embryo culture media lacking amino acids, such as human tubal fluid (HTF)50, and Earle’s51, high concentrations of glucose have been shown to impair human embryo development in culture52,53. An explanation for this observation has been derived from work on embryos from other mammalian species. Although glucose is not the preferred energy substrate of 2the cleavage-stage embryo, its uptake and utilization in a medium lacking amino acids is driven by its concentration in the culture medium54. As glycolysis is far less efficient in terms of energy production than oxidation, it is proposed that the premature utilization of glucose via glycolysis will culminate in inadequate energy production and hence impaired embryo development. This phenomenon is similar to that reported for certain types of tumor cell metabolism, and is known as the Crabtree effect55. The Crabtree effect, as described in tumor cells in culture, depends on the continued activity of hexokinase in the presence of increasing product, glucose-6-phosphate. The isoform of hexokinase present in these cells must therefore be a form of the enzyme that is not completely inhibited by glucose-6-phosphate. Kinetic analysis of hexokinase in preimplantation mouse embryos has revealed that there is a switch from isoform I at the zygote to isoform II at the blastocyst stage56. Indeed, these two isoforms have differing sensitivities to phosphate, with the inhibition of isoform I by glucose-6-phosphate being overcome by phosphate57, thereby helping to explain the susceptibility of the cleavagestage embryo to inhibition by glucose. As a direct consequence of the inhibitory nature of glucose under certain culture conditions, there has been a trend to remove glucose from embryo culture media53,58. However, it is important to consider that glucose is actually present in the female reproductive tract4, and that the oocyte and embryo possess a carrier mechanism for glucose entry into the cell42–45. Therefore, the inhibitory action of glucose in simple culture media should at best be considered as an in vitro-induced artifact. Importantly, the impairment of metabolic function in the embryo by glucose is not evident when specific

Metabolic requirements during preimplantation development

41

amino acids50 and the chelator ethylenediaminetetraacetic acid (EDTA) are present in the culture medium3. When zygotes of CF1 mice are cultured in conventional culture media, which contain glucose but lack amino acids and EDTA, they arrest in development at the 2-cell stage. Such arrested embryos exhibit a significant increase in glycolysis with a concomitant decrease in pyruvate oxidation13. The addition of amino acids and EDTA to the culture medium alleviates developmental arrest at the 2-cell stage and facilitates growth to the blastocyst59. Analysis of embryo metabolism revealed that amino acids and EDTA act through independent mechanisms to suppress glycolytic activity in the early embryo while increasing the levels of oxidation within the embryo1,13,60 (Figure 2.1a). Furthermore, amino acids and EDTA act in synergy to minimize the adverse metabolic effect of glucose on the cleavage-stage embryo, which is associated with an improvement in embryo development. Importantly, when zygotes from F1 mice were cultured to the 2cell stage under the same conditions as those used for the CF1-derived embryos, their metabolism was unaffected by culture conditions (Figure 2.1b). This therefore indicates that there are inherent differences between the embryos of different mouse strains in their ability to regulate their energy metabolism in culture, and that this in turn is related to their ability to develop in vitro. Indeed it does appear that 2cell embryos from F1 mice can maintain a high ATP/ADP ratio in culture, thereby minimizing glucose flux through glycolysis1. In contrast, 2-cell embryos from CF1 mice cannot maintain a high ATP/ADP ratio in culture conditions which are associated with developmental arrest, i.e. medium containing glucose but lacking amino acids and EDTA. As a result of the fall in the ATP/ADP ratio in CF1 embryos, glycolytic activity increases (Figure 2.2)61. Thus, it would appear that high levels of glycolysis in the cleavage-stage embryo are not consistent with embryo development in culture, and that suppression of glycolysis culminates in increased embryo development. In further support of this hypothesis it has been shown that, under conditions that would otherwise result in developmental arrest, the inclusion of a specific inhibitor of the glycolytic enzyme phosphoglycerate kinase, cibacron blue, facilitates development of CF1 zygotes to the 2-cell stage60. So under the appropriate, more physiological, culture conditions glucose need not be removed from the medium. In light of its roles other than as an energy source (see below), it would appear prudent to keep glucose in the culture medium at low levels initially, and then at increasing concentrations from the 8-cell stage onwards. Certainly, after the 8-cell stage the presence of glucose appears to be critical59.

Significance of glucose to the embryo The high levels of glycolysis exhibited by the mammalian blastocyst have been interpreted as the embryo’s adaptation to its imminent invasion of the endometrium, which through histology has been shown to remain avascular for a period of up to 12 h, and will therefore be relatively anoxic62,63. Subsequently, glycolysis will be the sole means of generating energy at this time. However, this may not be the sole explanation for the high levels of glycolysis in the blastocyst. An alternative explanation for the metabolism of glucose by the blastocyst is that as well as being used to generate energy for blastocoel expansion and mitosis, glucose will be required for the synthesis of

An atlas of human blastocysts 42

triacylglycerols and phospholipids, and as a precursor for complex sugars of mucopolysaccharides and glycoproteins. All of these are required by rapidly dividing cells64–66. Glucose metabolized by the pentose phosphate pathway (PPP) generates ribose moieties required for nucleic acid synthesis, and the NADPH (reduced nicotinamideadenine dinucleotide phosphate) required for the biosynthesis of lipids and other complex molecules65,67. NADPH is also required for the reduction of intracellular glutathione, an important antioxidant for the embryo68. The synthesis of nucleic acids is therefore an important biosynthetic role for glucose in the blastocyst. It has been proposed that high levels of aerobic glycolysis, such as that observed in the mammalian blastocyst, will ensure that there is sufficient substrate available for biosynthetic pathways, such as DNA replication, RNA transcription and synthesis of new membranes, at the required times during cellular proliferation69,70. This in turn suggests that there are times within the cell cycle during which the PPP is more active then others. Interestingly, the work of Hewitson and Leese71 indicates that the inner cell mass (ICM) generates its energy predominantly from glycolysis. An adequate supply of glucose may therefore be important for the optimal development of the ICM. In support of this, when mouse zygotes were cultured to the blastocyst in the absence of glucose, although resultant blastocysts implanted in the uterus after transfer, significantly more fetuses were lost compared with the control blastocysts which had developed in the presence of glucose59, indicating impaired ICM development or function.

Significance of pyruvate and lactate to the embryo Pyruvate enters the embryo both passively and by means of a facilitated carrier28,44, and is the preferred nutrient of the cleavage stage embryo of several species, including the human5. Although lactate is readily taken up, and can be metabolized to some degree, it cannot support the first cleavage division in the mouse18. Inside the embryo, pyruvate and lactate are converted by the enzyme lactate dehydrogenase (LDH) through the following reaction:

A primary function of pyruvate and lactate in cells is to regenerate NAD+ (oxidized nicotinamide-adenine dinucleotide) for subsequent use in glycolysis when under anaerobic conditions, and for the embryo this is of greatest significance at the blastocyst stage. Cytosolic regeneration of NAD+ is required, as the cytoplasmic and mitochondrial pools of NADH (reduced NAD+) are not shared. Rather, the reducing power between these two distinct cellular compartments is transferred through a specific system such as the malate/aspartate shuttle. It has been demonstrated that the malate/aspartate shuttle is involved in the metabolism of carboxylic acids and certain amino acids in the mouse zygote and cleavage-stage embryo. Significantly, a reduction of activity of this shuttle at the blastocyst stage may be responsible for aberrant levels of lactate production by blastocysts developed in vitro72,73.

Metabolic requirements during preimplantation development

43

It has been shown that the mouse zygote and blastocyst differ in their ability to metabolize pyruvate and lactate, and that such differences can only be accounted for by a change in the intracellular NAD/NADH ratio, which in turn is affected by the ratio of pyruvate/lactate74. Therefore, by changing the ratio of certain medium components one can inadvertently change the ratio of important intracellular regulators. For example, changing the concentration of lactate in the culture medium can have a significant effect on mouse embryo viability75. This effect is stage-specific, with different stages of development having different requirements for lactate to maintain viability75.

Different requirements of blastomeres after differentiation As discussed, low levels of glycolysis are consistent with development of the cleavagestage embryo in culture. However, as the later-stage embryo (morula and blastocyst) has a more glucose-dependent metabolism, this implies that different culture conditions will be required to support the changing metabolism of the embryo. For example, conditions which favor the development of the zygote, such as the limitation of glucose or the inhibition of a glycolytic enzyme (for example by EDTA), may well interfere with the development of the ICM, thereby affecting subsequent fetal development. Indeed this does appear to be the case. If mouse embryos are cultured to the blastocyst stage in the continual presence of EDTA, which inhibits glycolysis, then resultant fetal development is significantly lower than for those blastocysts which have been exposed to EDTA only for the first 48 h of culture from the zygote59. Similarly, the presence of EDTA in the culture medium for the first 72 h of culture significantly improved development of cattle embryos to the 8-cell stage. However, if the resultant 8-cell embryos were left in the presence of EDTA then the resultant blastocysts had a significantly smaller ICM, while the number of cells in the trophectoderm was unaffected76 (Figure 2.3). Such data highlight the differing metabolism of the cleavage-stage and post-compaction embryo, and further support the hypothesis that different culture conditions are required at different stages of development in order to satisfy the changing requirements of the embryo77–79.

‘Metabolic control’ hypothesis In light of the inherent susceptibility of the mammalian embryo to culture-induced stress in vitro, the ‘metabolic control’ hypothesis has been developed1. Under suboptimal culture conditions the embryo exhibits abnormal energy metabolism, or expressed another way the embryo undergoes metabolic transformation. It is proposed that the ability of a given embryo to regulate against such metabolic transformation is related to its ability to develop in culture and ultimately to develop into a viable fetus after transfer. Interestingly, metabolic transformation is not restricted to the cleavage-stage embryo, but is also manifest at the blastocyst stage too. Menke and McLaren80 were the first to report that mouse blastocysts cultured from the 8-cell stage in a balanced salt solution with added carbohydrates had an impaired oxidative ability, compared with blastocysts developed in vivo. Subsequently, Gardner and Leese27 reported that culturing mouse

An atlas of human blastocysts 44

morulae for around 12 h in medium MTF (mouse tubal fluid: balanced salt solution with carbohydrates but lacking amino acids) culminated in the production of blastocysts which exhibited very high levels of lactate production, inferring impaired oxidative capacity. Whereas a mouse blastocyst developed in vivo will convert around 40–50% of glucose consumed to lactate26,27, those blastocysts obtained from cultured morulae converted almost 100% of the glucose consumed to lactate. Subsequently Gardner and Sakkas75 found that supplementing a culture medium based upon a simple balanced salt solution with amino acids and vitamins abolished this metabolic transformation. Lane and Gardner81 then went on to investigate the timing of such metabolic transformation in the mouse blastocyst. It was observed that exposure to culture medium MTF for just 3 h resulted in an increase in lactate production, with maximum perturbation occurring after 6 h. Interestingly, the rate of glucose uptake was not affected by the incubation conditions, but rather its fate (Figure 2.4). Furthermore, the ability of blastocysts to oxidize pyruvate was significantly impaired after 6 h of culture in MTF. The addition of amino acids and vitamins to MTF independently decreased the excessive lactate production of blastocysts incubated for 6 h, and acted in synergy to reduce lactate production further. Moreover, the presence of both amino acids and vitamins in the incubation medium maintained the ability of the blastocyst to oxidize pyruvate. Perhaps the more important finding from this work was that the impaired metabolism observed after just 6 h of culture was associated with a significant reduction in blastocyst implantation and subsequent fetal development after transfer, compared with blastocysts flushed from the uterus and transferred to a recipient, or compared with blastocysts maintained in culture for 6 h but in the presence of both amino acids and vitamins. Clearly then, metabolic transformation is associated with loss of viability. This therefore leads to the question whether assessment of an embryo’s metabolism can be used to determine its viability before transfer, i.e. to select the most viable embryos from a given cohort.

Assessment of viability using metabolic criteria Owing to the limited amount of material available for study of the preimplantation mammalian embryo, miniaturizations of standard methods of biochemical analysis have to be employed. Two approaches have been used with great success: radiolabelled substrates in microincubation chambers, and ultramicrofluorescence. However, for the assessment of embryo viability prior to transfer, the former approach is not a realistic proposition, and so the latter approach is considered in detail here. Ultramicrofluorescence was developed by Mroz and Lechene and co-workers35. This technique was subsequently adapted37,38 to determine the nutrient uptake of an individual embryo at each successive stage of development in culture. Ultramicrofluorescence is a totally non invasive approach, and is based upon placing an individual embryo in a known volume of defined culture medium and then removing serial samples of culture medium from around the embryo and analyzing them for nutrient composition (Figure 2.5). The volumes of culture medium removed can be as small as nano- or picoliters. Such small volumes are manipulated with specially constructed micropipettes manufactured on

Metabolic requirements during preimplantation development

45

a microforge and calibrated with tritiated water. These pipettes are manipulated using a micromanipulator, and the fluids taken up and expelled using an air-filled syringe. The sub-microliter samples of fluid are then analyzed fluorometrically (Figure 2.6) using a fluorescence microscope with photometry attachments. In 1980, Renard and colleagues86 observed that day-10 cattle blastocysts which had a glucose uptake higher than 5 µg/h developed better both in culture and in vivo after transfer than those blastocysts with a glucose uptake below this value. However, owing to the insensitivity of the spectrophotmetric method employed, they could not quantitate glucose uptake by younger embryos. Rieger87 subsequently demonstrated that morphologically normal day-7 cattle blastocysts took up significantly more radiolabelled glucose than degenerating ones. In 1987, using the relatively new technique of noninvasive microfluorescence, Gardner and Leese88 mea-sured glucose uptake by individual day-4 mouse blastocysts prior to transfer to recipient females. Those embryos that went to term had a significantly higher glu-cose uptake in culture than those embryos that failed to develop after transfer. However, such studies were retro-spective, and as such could not conclusively demonstrate whether it was possible to identify viable embryos prior to transfer using metabolism as a marker. Subsequently, a study of the metabolism of day-7 cattle blastocysts before and after cryopreservation showed that it was possible to identify those blastocysts capable of re-expansion in the hours immediately postthaw89. Those blastocysts that survived the freeze-thaw procedure had a significantly higher glucose uptake and lactate production than those embryos that did not re-expand and subsequently died (Figure 2.7)89. Of greater significance, however, was the observation that there was no overlap in the distribution of glucose uptake by the viable and non-viable embryos, and very little overlap of lactate production, suggesting that it may therefore be possible to use metabolic criteria for prospective selection of viable embryos89. Interestingly pyruvate uptake by blastocysts did not reflect their subsequent viability. Therefore, in a prospective study, Lane and Gardner90 used glucose uptake and lactate production to determine glycolytic activity in individual day-5 mouse blastocysts prior to transfer. Consistent with the metabolic control hypothesis, blastocysts were classified as viable or non-viable according to their rate of glucose uptake and lactate production. Mouse blastocysts of equal dimensions and morphology subsequently had their metabolism quantitated non-invasively and were then classified as either viable or nonviable. It was found that those blastocysts which exhibited a pattern of glycolytic utilization similar to that of embryos developed in vivo had a developmental potential of 80%, while those blastocysts which exhibited an excessive lactate production (i.e. aberrant glycolytic activity) had a developmental potential of only 6% (Figure 2.8). Interestingly, when a retrospective analysis of glucose uptakes was performed on the blastocysts transferred in this study, those blastocysts classified as viable has a significantly higher rate of glucose uptake than those blastocysts classified as non-viable. Therefore, it would appear that both the rate of nutrient uptake and its subsequent fate are important determinants of embryo viability. Importantly, studies of nutrient uptake and subsequent viability have been performed on the human embryo. In a retrospective analysis, Conaghan and colleagues91 observed an inverse relationship between pyruvate uptake by 2–8-cell embryos and subsequent pregnancy. However, it is important to note that such measurements were performed

An atlas of human blastocysts 46

prior to human embryonic genome activation. It is therefore plausible that the observed differences in pyruvate uptake reported by Conaghan and colleagues91 reflected differences inherited from the oocyte, and did not represent the true physiology of the later-stage embryo. Furthermore, the medium used to assess nutrient consumption was a simple one, lacking amino acids and vitamins. In a study of human morulae and blastocysts of different degrees of expansion, no conclusive data were generated on the ability of nutrient consumption or utilization to predict pregnancy outcome92. Again, however, the medium used to assess embryo metabolism was a simple one, lacking pyruvate, lactate, amino acids and vitamins. Under such severe culture conditions, the resultant stress on the embryos would have been enormous, and therefore it is questionable whether any meaningful data could have been obtained. In fact, one would expect embryos undergoing such a treatment to be compromised. In contrast, Van den Bergh and associates93 showed that, in patients who conceived following blastocyst transfer, embryos had a significantly lower glycolytic activity than those embryos which did not establish a pregnancy. Significantly, in the work of Van den Bergh and associates93, a complete medium was used for the metabolic assessment, thereby alleviating the culture-induced stress associated with the work of Jones and colleagues92. More recently, two studies have determined the relationship between embryo nutrition and subsequent development in vitro94,95. Gardner and colleagues94 determined that glucose consumption on day 4 by human embryos was twice as high in those embryos that went on to form blastocysts. Furthermore, it was determined that blastocyst quality affected glucose uptake. Poor-quality blastocysts consumed significantly less glucose than top-scoring embryos. Significantly, within a cohort of human blastocysts from the same patient with the same alpha-numeric score, i.e. 4AA, there existed a significant spread of metabolic activities. These embryos were cultured in sequential media G1 and G2, and their metabolism assessed in medium G2 in order to prevent metabolic transformation. Therefore, assessing metabolic activity and metabolic normality may prove to be a feasible way of determining embryonic ‘health’ (Figure 2.9). In a study of amino acids, Houghton and co-workers95 determined that alanine release into the surrounding medium on day 2 and day 3 was highest in those embryos that did not form blastocysts.

Culture of the human embryo Table 2.4 gives the compositions of the sequential media G1.2 and G2.2. The media were developed after taking into account the data described in this chapter. In G1.2, carbohydrates are present at concentrations measured in the human Fallopian tube at the time when the embryo

Metabolic requirements during preimplantation development

47

Table 2.4 Compositions of sequential culture media G1.2 and G2.2 Component

mmol/l

Component

mmol/l

G1.2 (cleavage-stage development) Sodium chloride

90.08 alanyl-glutamine

0.5

Potassium chloride

5.5 alanine

0.1

Sodium phosphate

0.25 aspartate

0.1

Magnesium sulfate Bicarbonate Calcium chloride

1.0 asparagine

0.1

25.0 glutamate

0.1

1.8 glycine

0.1

proline

0.1

Glucose

0.5 serine

0.1

Lactate

10.5 taurine

0.1

Pyruvate

0.32 EDTA

0.01

90.08 arginine

0.6

Potassium chloride

5.5 cystine

0.1

Sodium phosphate

0.25 histidine

0.2

G2.2 (blastocyst development) Sodium chloride

Magnesium sulfate Bicarbonate Calcium chloride

1.0 isoleucine

0.4

25.0 leucine

0.4

1.8 lysine

0.4

methionine

0.1

Glucose

3.15 phenylalanine

0.2

Lactate

5.87 threonine

0.4

Pyruvate

0.10 tryptophan

0.5

tyrosine

0.2

Alanyl-glutamine

1.0 valine

0.4

Alanine

0.1

Aspartate

0.1 choline chloride

0.0072

Asparagine

0.1 folic acid

0.0023

Glutamate

0.1 inositol

Glycine

0.1 nicotinamide

0.01 0.0082

An atlas of human blastocysts 48

Proline

0.1 pantothenate

0.0042

Serine

0.1 pyridoxal

0.0049

riboflavin

0.00027

EDTA, ethylenediaminetetraacetic acid

is present. In contrast, in medium G2.2, concentrations of carbohydrates are those found in the human uterus. The amino acids present in medium G1.2 are those present at high levels in mammalian oviduct fluids, and have been shown to stimulate the cleavage-stage embryo96,97 and minimize intracellular stress1,15,13,98, while those in G2.2 are those needed for the development of both the ICM and trophectoderm, which have different nitrogen requirements8. EDTA is present in medium G1.2 for the reasons listed above, while it is absent from medium G2.2, to ensure optimal development of the ICM. Vitamins are absent from medium G1.2 but present in G2.2, to facilitate sufficient oxidation in the blastocyst1,81. Using such sequential media it is possible to culture the human embryo, along with those of several other species97, to the blastocyst stage at high rates, culminating in a high implantation rate following transfer. Table 2.5 lists results of using these media in a clinical setting. Figure 2.10 shows the typical morphology of human blastocysts cultured in these sequential media. As discussed, amino acids are among the most important regulators of mammalian embryo development in culture1,3,97,102. However, amino acids both are metabolized by the embryos and spontaneously break down at 37°C to produce ammonium84, which impairs embryo development, ICM formation and fetal development84,103,104. The majority of ammonium released into the culture system comes from their spontaneous breakdown while at 37°C, which has profound implications for the storage and use of media containing them. Ideally, embryo culture media should be set up at least 4 h and no more than 18 h before use105, and such media should be renewed at least every 48 h84. Of the amino acids present in culture media, glutamine is known to be the most labile, and contributes a disproportionate amount of ammonium to the system84. This particular problem can be alleviated by the substitution of glutamine with alanyl-glutamine106, which does not readily deaminate at 37°C. More recently, the concept of using sequential media has been challenged, and the use of one medium (KSOMAA) to support all the preimplantation stages has been proposed as an effective culture method107. It is evident that one culture medium will support blastocyst development in a number of mammalian species including the human108. However, as discussed at length above, there is considerable difference in the ability to culture blastocysts and the ability to culture viable blastocysts78,79.The question is, therefore, can one medium support the development of blastocysts with the same viability as those blastocysts developed in sequential media? The answer to this question, at least in animal models, is no 97. The medium KSOMAA by default does not allow for any nutrient gradients, and therefore will induce some form of metabolic stress on the embryo as described in detail above. Furthermore, the medium KSOMAA contains EDTA. Subsequently, the embryo will be exposed to EDTA post-compaction, and this will have a negative effect on ICM development as discussed. Analysis of mouse and cow embryos cultured in either sequential media G1/G2 or KSOMAA have revealed that those embryos

Metabolic requirements during preimplantation development

49

cultured in KSOMAA do indeed have reduced embryo development, reduced blastocyst cell number, impaired ICM development and lower viability after transfer97. These findings are consistent with the data presented in this chapter on nutrient requirements and metabolism of the embryo. Significantly, should KSOMAA be used, it is necessary to renew the medium every 48 h as it contains 1 mmol/l glutamine, and produces embryo toxic levels of ammonium within the duration of the culture period. In an ideal world, rather than simply using two culture media in sequence, the embryo in culture could be exposed to any number of nutrient gradients, obtainable through a perfusion system10. Figure 2.11 shows the basic concept of such a system. Recently, with the introduction of microfluidic cells, the concept of perfusion culture in embryology is becoming a reality109,110. Embryo culture systems could be about to change in the clinical in vitro fertilization (IVF) laboratory. Although this chapter endeavors to place the nutrition and metabolism of the embryo into a meaningful context, it would be remiss not to discuss, albeit briefly, the significance of macromolecules in the development of the embryo. Relative to other culture medium components, macromolecules have received relatively little attention. Recently, however, studies have demonstrated the benefit of the inclusion of the glycosaminoglycan hyaluronan in culture media on embryo development111, cryosurvival112–114 and transfer outcome111,115. Furthermore, recombinant albumin is now available, and not only has it been shown to be equally as effective in an IVF/embryo culture system as blood-derived human serum albumin114,116–122, but it also increases embryo cyrosurvivability113. Significantly, the introduction of genetically engineered macromolecules

Table 2.5 Summary of blastocyst transfer data at the Colorado Center for Reproductive Medicine. From reference 99, with permission IVF patients*

Oocyte donors

Number of patients

401

211

Number of patients having embryo transfer

395

211

Mean age (±SEM) in years

33.4±0.2

40.6±0.3 recipients

Age range (years)

20–43

27–50

FSH (IU/1) (mean±SEM)

6.7±0.1

6.1±0.1 donors

Patients with ICSI (%)

40.4

39.3

Number of pronucleate embryos (mean±SEM)

14.5±0.3

15.2±0.4

Blastocyst development on day 5 (%)

44.1

51.7

Blastocyst development on day 6 (%)

8.1

8.3

Total blastocyst development (%)

52.2

60.0

Number of embryos transferred (mean±SEM)

2.2±0.03

2.1±0.03

Patients with embryo freezing (%)†

75.3

85.3

An atlas of human blastocysts 50

Mean number of blastocysts frozen (mean±SEM)

4.3±0.2

5.6±0.3

Implantation rate (fetal sac) (%)‡

50.1

62.1

Implantation rate (fetal heart) (%)‡

46.4

60.8

Clinical pregnancy rate (%)**

68.6

79.6 †

*In vitro fertilization (IVF) patients had at least ten follicles; only blastocysts scoring 3BB or higher by the afternoon of day 6 were cryopreserved; ‡implantation rates are expressed as fetal sac or heart/blastocyst transferred. The calculations included every patient who had an embryo transfer and not just those who subsequently became pregnant; **includes six patients in the blastocyst culture group who did not have an embryo transferred on day 5 owing to embryonic arrest at the cleavage stage. Clinical pregnancy was determined by the presence of a fetal heart beat; FSH, follicle stimulating hormone; ICSI, intracytoplasmic sperm injection

means that the potential transmission of blood-borne pathogens such as human immunodeficiency virus (HIV) and prions is effectively eliminated. Furthermore, the lotto-lot variation associated with serum derived albumin119,120 is also eliminated. Such changes have resulted in the development of new culture systems97. The adverse effects of whole serum on embryo development, metabolism and ultrastructure have been well documented. The use of serum in a clinical embryo culture system cannot be considered morally acceptable.

Conclusions It is evident that suboptimal culture conditions induce a considerable degree of cellular stress in the preimplantation embryo. A key manifestation of such culture-induced stress is abnormal energy metabolism. Typically, under suboptimal conditions, glycolytic activity increases at the expense of oxidation, culminating in inadequate energy production and impaired embryo development. Culture conditions which increase embryo development in vitro appear to help the embryo maintain a more in vivo-like metabolism, i.e. low levels of glycolysis prior to compaction, while maintaining the embryo’s ability to oxidize a percentage of the glucose consumed at the blastocyst stage. The normalcy of metabolic activity of an embryo appears to be related to both developmental capacity in culture and subsequent viability after transfer. Therefore, the non-invasive assessment of metabolism of an individual embryo within a given cohort prior to transfer may help identify those embryos most likely to give rise to a successful pregnancy. By meeting the nutritional requirements of the embryo as it develops and differentiates (including carbohydrate and amino acid gradients), and by reducing stress, one can support the development of highly viable blastocysts in culture. By using sequential media it is now possible to obtain in vivo rates of embryo development in vitro in animal models97,121.

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51

References 1. Gardner DK. Changes in requirements and utilization of nutrients during mammalian preimplantation embryo development and their significance in embryo culture. Theriogenology 1998; 49:83–102 2. Lane M, Gardner DK. Regulation of ionic homeostasis by mammalian embryos. Semin Reprod Med 2000; 18:195–204 3. Gardner DK, Pool TB, Lane M. Embryo nutrition and energy metabolism and its relationship to embryo growth, differentiation, and viability. Semin Reprod Med 2000; 18:205–18 4. Gardner DK, Lane M, Calderon I, Leeton J. Environment of the preimplantation human embryo in vivo: metabolite analysis of oviduct and uterine fluids and metabolism of cumulus cells. Fertil Steril 1996; 65:349–53 5. Hardy K, Hooper MA, Handyside AH, Rutherford AJ, Winston RM, Leese HJ. Non-invasive measurement of glucose and pyruvate uptake by individual human oocytes and preimplantation embryos. Hum Reprod 1989; 4:188–91 6. Gott AL, Hardy K, Winston RM, Leese HJ. Non-invasive measurement of pyruvate and glucose uptake and lactate production by single human preimplantation embryos. Hum Reprod 1990; 5:104–8 7. Gardner DK, Lane M, Stevens J, Schoolcraft WB. Noninvasive assessment of human embryo nutrient consumption as a measure of developmental potential. Fertil Steril 2001; 76:1175–80 8. Lane M, Gardner DK. Differential regulation of mouse embryo development and viability by amino acids. J Reprod Fertil 1997; 109:153–64 9. Steeves TE, Gardner DK. Temporal and differential effects of amino acids on bovine embryo development in culture. Biol Reprod 1999; 61:731–40 10. Gardner DK. Mammalian embryo culture in the absence of serum or somatic cell support. Cell Biol Int 1994; 18:1163–79 11. Crosby IM, Gandolfi F, Moor RM. Control of protein synthesis during early cleavage of sheep embryos. J Reprod Fertil 1988; 82:769–75 12. Rieger D, Loskutoff NM, Betteridge KJ. Developmentally related changes in the metabolism of glucose and glutamine by cattle embryos produced and co-cultured in vitro. J Reprod Fertil 1992; 95:585–95 13. Gardner DK, Lane M. The 2-cell block in CF1 mouse embryos is associated with an increase in glycolysis and a decrease in tricarboxylic acid (TCA) cycle activity: alleviation of the 2-cell block is assocated with the restoration of in vivo metabolic pathway activities. Biol Reprod 1993; 48(Suppl 1):152 14. Van Winkle LJ, Haghighat N, Campione AL. Glycine protects preimplantation mouse conceptuses from a detrimental effect on development of the inorganic ions in oviductal fluid. J Exp Zool 1990; 253:215–19 15. Edwards LJ, Williams DA, Gardner DK. Intracellular pH of the mouse preimplantation embryo: amino acids act as buffers of intracellular pH. Hum Reprod 1998; 13:3441–8 16. Liu Z, Foote RH. Development of bovine embryos in KSOM with added superoxide dismutase and taurine and with five and twenty percent O2. Biol Reprod 1995; 53:786–90 17. Lindenbaum A. A survey of naturally occurring chelating ligands. Adv Exp Med Biol 1973; 40:67–77 18. Biggers JD, Whittingham DG, Donahue RP. The pattern of energy metabolism in the mouse oocyte and zygote. Proc Natl Acad Sci USA 1967; 58:560–7 19. Biggers JD, Gardner DK, Leese HJ. Control of carbohydrate metabolism in preimplantation mammalian embryos. In Rosenblum IY, Heyner S, eds. Growth Factors in Mammalian Development. Boca Raton: CRC Press, 1989:19–32 20. Leese HJ. Metabolism of the preimplantation mammalian embryo. Oxf Rev Reprod Biol 1991; 13:35–72

An atlas of human blastocysts 52 21. Mills RM, Brinster RL. Oxygen consumption of preimplantation mouse embryos. Exp Cell Res 1967; 47:337–44 22. Houghton FD, Thompson JG, Kennedy CJ, Leese HJ. Oxygen consumption and energy metabolism of the early mouse embryo. Mol Reprod Dev 1996; 44:476–85 23. Thompson JG, Partridge RJ, Houghton FD, Cox Cl, Leese HJ. Oxygen uptake and carbohydrate metabolism by in vitro derived bovine embryos. J Reprod Fertil 1996; 106:299–306 24. Brinster RL. Studies on the development of mouse embryos in vitro. IV. Interaction of energy sources. J Reprod Fertil 1965; 10:227–40 25. Brinster RL, Thomson JL. Development of eight-cell mouse embryos in vitro. Exp Cell Res 1966; 42:308–15 26. Wales RG. Measurement of metabolic turnover in single mouse embryos. J Reprod Fertil 1986; 76:717–25 27. Gardner DK, Leese HJ. Concentrations of nutrients in mouse oviduct fluid and their effects on embryo development and metabolism in vitro. J Reprod Fertil 1990; 88:361–8 28. Leese HJ, Barton AM. Production of pyruvate by isolated mouse cumulus cells. J Exp Zool 1985; 234:231–6 29. Barbehenn EK, Wales RG, Lowry OH. The explanation for the blockade of glycolysis in early mouse embryos. Proc Natl Acad Sci USA 1974; 71:1056–60 30. Barbehenn EK, Wales RG, Lowry OH. Measurement of metabolites in single preimplantation embryos; a new means to study metabolic control in early embryos. J Embryol Exp Morphol 1978; 43:29–46 31. Telford NA, Watson AJ, Schultz GA. Transition from maternal to embryonic control in early mammalian development: a comparison of several species. Mol Reprod Dev 1990; 26:90–100 32. Epstein CJ, Smith SA. Amino acid uptake and protein synthesis in preimplantation mouse embryos. Dev Biol 1973; 33:171–84 33. Benos D, Biggers JD. Blastocyst fluid formation. In Mastroianni LJ, Biggers JD, eds. Fertilization and Embryonic Development In Vitro. New York: Plenum Press, 1981:283–97 34. Biggers JD, Bell JE, Benos DJ. Mammalian blastocyst: transport functions in a developing epithelium. Am J Physiol 1988; 255:C419–32 35. Leese HJ, Biggers JD, Mroz EA, Lechene C. Nucleotides in a single mammalian ovum or preimplantation embryo. Anal Biochem 1984; 140:443–8 36. Rozell MD, Williams JE, Butler JE. Changes in concentration of adenosine triphosphate and adenosine diphosphate in individual preimplantation sheep embryos. Biol Reprod 1992; 47:866– 70 37. Gardner DK, Leese HJ. Non-invasive measurement of nutrient uptake by single cultured preimplantation mouse embryos. Hum Reprod 1986; 1:25–7 38. Gardner DK, Lane M, Batt P. Uptake and metabolism of pyruvate and glucose by individual sheep preattachment embryos developed in vivo. Mol Reprod Dev 1993; 36:313–19 39. Leese HJ, Barton AM. Pyruvate and glucose uptake by mouse ova and preimplantation embryos. J Reprod Fertil 1984; 72:9–13 40. Thompson JG, Simpson AC, Pugh PA, Tervit HR. Requirement for glucose during in vitro culture of sheep preimplantation embryos. Mol Reprod Dev 1992; 31:253–7 41. Scott LA. Oocyte and embryo culture. In Keel BA, May JV, De Jonge CJ, eds. Handbook of the Assisted Reproduction Laboratory. Boca Raton: CRC Press, 2000; 197–219 42. Aghayan M, Rao LV, Smith RM, et al. Developmental expression and cellular localization of glucose transporter molecules during mouse preimplantation development. Development 1992; 115:305–12 43. Dan-Goor M, Sasson S, Davarashvili A, Almagor M. Expression of glucose transporter and glucose uptake in human oocytes and preimplantation embryos. Hum Reprod 1997; 12:2508–10 44. Gardner DK, Leese HJ. The role of glucose and pyruvate transport in regulating nutrient utilization by preimplantation mouse embryos. Development 1988; 104:423–9

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45. Hogan A, Heyner S, Charron MJ, et al. Glucose transporter gene expression in early mouse embryos. Development 1991; 113:363–72 46. Biggers JD, Stern S. Metabolism of the preimplantation mammalian embryo. Adv Reprod Physiol 1973; 6:1–59 47. Martin KL, Hardy K, Winston RM, Leese HJ. Activity of enzymes of energy metabolism in single human preimplantation embryos. J Reprod Fertil 1993; 99:259–66 48. Auerbach S, Brinster RL. Lactate dehydrogenase isozymes in mouse blastocyst cultures. Exp Cell Res 1968:53:313–15 49. Edwards LE, Gardner DK. Characterization of hexokinase kinetics in the preimplantation mouse embryo. Proc Fertil Soc Aust 1995; 14:28 50. Quinn P, Kerin JF, Warnes GM. Improved pregnancy rate in human in vitro fertilization with the use of a medium based on the composition of human tubal fluid. Fertil Steril 1985; 44:493– 8 51. Edwards RG. Test-tube babies. Nature (London) 1981; 293:253–6 52. Conaghan J, Handyside AH, Winston RM, Leese HJ. Effects of pyruvate and glucose on the development of human preimplantation embryos in vitro. J Reprod Fertil 1993; 99:87–95 53. Quinn P. Enhanced results in mouse and human embryo culture using a modified human tubal fluid medium lacking glucose and phosphate. J Assist Reprod Genet 1995; 12:97–105 54. Vella P, Lane M, Gardner DK. Induction of glycolysis in the day-3 mouse embryo by glucose. Biol Reprod 1997; 57(Suppl 1):26 55. Koobs DH. Phospate mediation of the Crabtree and Pasteur effects. Science 1972; 178:127–33 56. Edwards LE, Gardner DK. Characterisation of hexokinase kinetics in the preimplantation mouse embryo. Proc Fertil Soc Aust 1995; 14:28 57. Wilson JE. Hexokinases. Rev Physiol Biochem Pharmacol 1995; 126:65–198 58. Pool TB, Atiee SH, Martin JE. Oocyte and embryo culture: basic concepts and recent advances. Infertil Reprod Med Clin North Am 1998; 9:181–203 59. Gardner DK, Lane M. Alleviation of the ‘2-cell block’ and development to the blastocyst of CF1 mouse embryos: role of amino acids, EDTA and physical parameters. Hum Reprod 1996; 11:2703–12 60. Lane M, Gardner DK. Inhibiting 3-phosphoglycerate kinase by EDTA stimulates the development of the cleavage stage mouse embryo. Mol Reprod Dev 2001; 60:233–40 61. Gardner DK, Lane M. Alleviation of the 2 cell block in CF1 mouse embryos is associated with an increase in the ATP:ADP ratio and subsequent inhibition of PFK. Biol Reprod 1997;57(Suppl 1):216 62. Rogers PW, Murphy CR, Gannon BJ. Absence of capillaries in the endometrium surrounding the implanting rat blastocyst. Micron 1982; 13:373–4 63. Rogers PW, Murphy CR, Gannon BJ. Changes in the spatial organization of the uterine vasculature during implantation in the rat. J Reprod Fertil 1982; 65:211–14 64. Hume DA, Weidemann MJ. Role and regulation of glucose metabolism in proliferating cells. J Natl Cancer Inst 979; 62:3–8 65. Morgan MJ, Faik P. Carbohydrate metabolism in cultured animal cells. Biosci Rep 1981; 1:669–86 66. Mandel LJ. Energy metabolism of cellular activation, growth, and transformation. Curr Top Memb Trans 1986; 27:261–91 67. Reitzer LJ, Wice BM, Kennell D. The pentose cycle: control and essential function in HeLa cell nucleic acid synthesis. J Biol Chem 1980; 255:5616–26 68. Rieger D. Relationship between energy metabolism and development of the early embryo. Theriogenology 1992; 37:75–93 69. Newsholme EA, Crabtree B, Ardawi MS. The role of high rates of glycolysis and glutamine utilization in rapidly dividing cells. Biosci Rep 1985; 5:393–400 70. Newsholme EA. Application of metabolic-control logic to the requirements for cell division. Biochem Soc Trans 1990; 18:78–80

An atlas of human blastocysts 54 71. Hewitson LC, Leese HJ. Energy metabolism of the trophectoderm and inner cell mass of the mouse blastocyst. J Exp Zool 1993; 267:337–43 72. Gardner DK, Pool TB, Lane M. Embryo nutrition and energy metabolism and its relationship to embryo growth, differentiation, and viability. Semin Reprod Med 2000; 18:205–18 73. Lane M, Gardner DK. Regulation of substrate utilization in mouse embryos by the malateaspartate shuttle. Biol Reprod 2000; 62(Suppl 1):371 74. Lane M, Gardner DK. Lactate regulates pyruvate uptake and metabolism in the preimplantation mouse embryo. Biol Reprod 2000; 62:16–22 75. Gardner DK, Sakkas D. Mouse embryo cleavage, metabolism and viability: role of medium composition. Hum Reprod 1993; 8:288–95 76. Gardner DK, Lane MW, Lane M. EDTA stimulates cleavage stage bovine embryo development in culture but inhibits blastocyst development and differentiation. Mol Reprod Dev 2000; 57:256–61 77. Gardner DK, Lane M. Embryo culture systems. In Gardner DK, Trounson AO, eds. Handbook of In Vitro Fertilization. Boca Raton: CRC Press, 1993:85–114 78. Gardner DK, Lane M. Culture and selection of viable blastocysts: a feasible proposition for human IVF? Hum Reprod Update 1997; 3:367–82 79. Gardner DK, Lane M. Culture of viable human blasto cysts in defined sequential serum-free media. Hum Reprod 1998; 13(Suppl 3):148–59 80. Menke TM, McLaren A. Mouse blastocysts grown in vivo and in vitro: carbon dioxide production and trophoblast outgrowth. J Reprod Fertil 1970; 23:117–27 81. Lane M, Gardner DK. Amino acids and vitamins prevent culture-induced metabolic perturbations and associated loss of viability of mouse blastocysts. Hum Reprod 1998; 13:991–7 82. Gardner DK, Leese HJ.Assessment of embryo metabolism and viability. In Trounson AO, Gardner DK, eds. Handbook of In Vitro Fertilization, 2nd edn. Boca Raton: CRC Press, 1999:347–72 83. Gardner DK, Clarke RN, Lechene CP, Biggers JD. Development of a noninvasive ultramicrofluorometric method for measuring net uptake of glutamine by single preimplantation mouse embryos. Gamete Res 1989; 24:427–38 84. Gardner DK, Lane M. Amino acids and ammonium regulate mouse embryo development in culture. Biol Reprod 1993; 48:377–85 85. Johnson SK, Jordan JE, Dean RG, Page RD. The quantification of bovine embryo viability using a bioluminescent assay for lactate dehydrogenase. Theriogenology 1991; 35:425–33 86. Renard JP, Philippon A, Menezo Y. In-vitro uptake of glucose by bovine blastocysts. J Reprod Fertil 1980; 58:161–4 87. Rieger D. The measurement of metabolic activity as an approach to evaluating viability and diagnosing sex in early embryos. Theriogenology 1984; 21:138–49 88. Gardner DK, Leese HJ. Assessment of embryo viability prior to transfer by the noninvasive measurement of glucose uptake. J Exp Zool 1987; 242:103–5 89. Gardner DK, Pawelczynski M, Trounson AO. Nutrient uptake and utilization can be used to select viable day 7 bovine blastocysts after cryopreservation. Mol Reprod Dev 1996; 44:472–5 90. Lane M, Gardner DK. Selection of viable mouse blastocysts prior to transfer using a metabolic criterion. Hum Reprod 1996; 11:1975–8 91. Conaghan J, Hardy K, Handyside AH, Winston RM, Leese HJ. Selection criteria for human embryo transfer: a comparison of pyruvate uptake and morphology. J Assist Reprod Genet 1993; 10:21–30 92. Jones G, Trounson A, Vella P, et al. Glucose metabolism of human morula and blastocyst stage embryos and its relationship to viability after transfer. Reprod Biomed Online 2001; 3:124–32 93. Van den Bergh M, Devreker F, Emiliani S, Englert Y. Glycolytic activity: a possible tool for human blastocyst selection. Reprod BioMed Online 2001; 3(Suppl 1):8 94. Gardner DK, Lane M, Stevens J, Schoolcraft WB. Noninvasive assessment of human embryo nutrient consumption as a measure of developmental potential. Fertil Steril 2001; 76:1175–80

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95. Houghton FD, Hawkhead JA, Humpherson PG, et al. Non-invasive amino acid turnover predicts human embryo developmental capacity. Hum Reprod 2002; 17:999–1005 96. Lane M, Gardner DK. Nonessential amino acids and glutamine decrease the time of the first three cleavage divisions and increase compaction of mouse zygotes in vitro. J Assist Reprod Genet 1997; 14:398–403 97. Gardner DK, Lane M. Development of viable mammalian embryos in vitro: evolution of sequential media. In Cibelli J, Lanza RP, Campbell KHS, West MD, eds. Principles of Cloning. San Diego: Academic Press, 2002:187–213 98. Lane M, Gardner DK. Regulation of ionic homeostasis by mammalian embryos. Semin Reprod Med 2000; 18:195–204 99. Gardner DK, Lane M, Schoolcraft WB. Physiology and culture of the human blastocyst. J Reprod Immunol 2002; 55:85–100 100. Gardner DK, Lane M, Stevens J, Schlenker T, Schoolcraft WB. Blastocyst score affects implantation and pregnancy outcome: towards a single blastocyst transfer. Fetil Steril 2000; 73:1155–8 101. Gardner DK, Schoolcraft WB. In-vitro culture of human blastocysts. In Jansen R, Mortimer D, eds. Towards Reproductive Certainty: Fertility and Genetics Beyond 1999. Carnforth, UK: Parthenon Publishing 1999:378–88 102. Gardner DK. Culture of mammalian embryos in the absense of serum and somatic cells. Cell Biol Int 1994; 18:1163–79 103. Lane M, Gardner DK. Culture of preimplantation mouse embryos in the presence of amino acids increases post implantation development whilst the concomitant production of ammonium induces birth defects. J Reprod Fertil 1994; 102:305–12 104. Lane M, Gardner DK. Ammonium affects ICM development, metabolism, intracellular pH, and fetal growth rates. Biol Reprod 2002; 66(Suppl 1):17 105. Gardner DK, Lane M. Embryo culture. In Gardner DK, Weissman A, Howles C, Shoham Z, eds. Textbook of Assisted Reproductive Techniques. London: Martin Dunitz, 2001:203–22 106. Gardner DK, Schoolcraft WB, Wagley L, Schlenker T, Stevens J, Hesla J. A prospective randomized trial of blastocyst culture and transfer in in-vitro fertilization. Hum Reprod 1998; 13:3434–40 107. Biggers JD, Racowsky C. The development of fertilized human ova to the blastocyst stage in KSOMAA medium: is a two-step protocol necessary? Reprod BioMed Online 2002; 5:133–40 108. Bolton VN, Wren ME, Parsons JH. Pregnancies after in vitro fertilization and transfer of human blastocysts. Fertil Steril 1991; 55:830–2 109. Beebe DJ, Moore JS, Yu Q, et al. Microfluidic tectonics: a comprehensive construction platform for microfluidic systems. Proc Natl Acad Sci USA 2000; 97:13488–93 110. Glasgow IK, Zeringue HC, Beebe DJ, et al. Handling individual mammalian embryos using microfluidics. IEEE Trans Biomed Eng 2001; 48:570–8 111. Gardner DK, Rodriegez-Martinez H, Lane M. Fetal development after transfer is increased by replacing protein with the glycosaminoglycan hyaluronan for mouse embryo culture and transfer. Hum Reprod 1999; 14:2575–80 112. Gardner DK, Maybach JM, Lane M. Hyaluronan and rHSA increase blastocyst cryosurvival. Presented at the 17th World Congress on Fertility and Sterility, Melbourne, Australia, November 2001:226 113. Stojkovic M, Kolle S, Pein S, et al. Effects of high concentrations of hyaluronan in culture medium on development and survival rates of fresh and frozen-thawed bovine embryos produced in vitro. Reproduction 2002; 124:141–53 114. Lane M, Maybach JM, Hooper K, Hasler JF, Gardner DK. Cryo-survival and development of bovine blastocysts are enhanced by culture with recombinant albumin and hyaluronan. Mol Reprod Dev 2003; 64:70–8

An atlas of human blastocysts 56 115. Schoolcraft WB, Lane M, Stevens J, Gardner DK. Increased hyaluronan concentration in the embryo transfer medium results in a significant increase in human embryo implantation rate. Fertil Steril 2002; 76 (Suppl 3):S5 116. Gardner DK, Lane M. Recombinant human serum albumin and hyaluronan can replace bloodderived albumin in embryo culture media. Fertil Steril 2000; 74 (Suppl 3):S31 117. Bungum M, Humaidan P, Bungum L. Recombinant human albumin as protein source in culture media used for IVF: a prospective randomized study. Reprod BioMed Online 2002; 4:233–6 118. Bavister BD, Kinsey DL, Lane M, Gardner DK. Recombinant human albumin supports hamster in vitro fertilization. Hum Reprod 2003; in press 119. Batt PA, Gardner DK, Cameron AW. Oxygen concentration and protein source affect the development of preimplantation goat embryos in vitro. Reprod Fertil Dev 1991; 3:601–7 120. McKiernan SH, Bavister BD. Different lots of bovine serum albumin inhibit or stimulate in vitro development of hamster embryos. In Vitro Cell Dev Biol 1992; 28A: 154–6 121. Reed LC, Lane M, Gardner DK. In vivo rates of mouse embryo development can be attained in vitro. Theriogenology 2003; in press

Figure 2.1 Glycolytic activity and pyruvate oxidation of CF1 (a) and F1 (b) 2-cell embryos cultured from the zygote. MTF, mouse tubal fluid medium; aa, amino acids. Significantly different from embryos in MTF:

Metabolic requirements during preimplantation development

*p<0.05; **p<0.01. Data from reference 13

Figure 2.2 Adenosine nucleotide ratio in CF1 and F1 2-cell embryos cultured from the zygote either under conditions which facilitated development through the 2-cell block (MTF+ ethylenediaminetetraacetic acid (EDTA)+amino acids), or under conditions which induced cleavage arrest at the 2-cell stage in CF1 embryos (MTF)59. Values are mean ± SEM of ten replicates. Open bars, MTF+EDTA+amino acids; closed bars, MTF. Like pairs are significantly different; a, p<0.01; b, p<0.05. Data from reference 61

57

An atlas of human blastocysts 58

Figure 2.3 Effect of EDTA present in culture medium on the development and differentiation of bovine zygotes to the blastocyst stage:−/− embryos cultured for 72 h in medium SOFaa (synthetic oviduct fluid with amino acids) lacking EDTA followed by culture in medium SOFaa lacking EDTA; +/− embryos cultured for 72 h in medium SOFaa with 100 µmol/l EDTA followed by culture in medium SOFaa lacking EDTA; +/+ embryos cultured for 72 h in medium SOFaa with 100 µmol/l EDTA followed by culture in medium SOFaa with 100 µmol/l EDTA. Open bars represent percentage morula/blastocyst development, dark bars represent total cell number, gray bars represent inner cell mass (ICM) cell number. Like pairs of letters are significantly different: a, h, p<0.05; b, c, d, e, f, g, i, p<0.01. Data from reference 76

Metabolic requirements during preimplantation development

Figure 2.4 Effect of amino acids and vitamins in modified mouse tubal fluid (MTF) medium on glucose uptake and lactate production by mouse blastocysts. MTF+AA: MTF supplemented with 20 amino acids; MTF+VIT: MTF supplemented with vitamins; MTF +AA+VIT: MTF supplemented with Eagle’s 20 amino acids and Eagle’s vitamins. Open bars represent glucose uptake. Closed bars represent lactate production. Significantly different from in vivodeveloped blastocysts: *p<0.05; **p<0.01. Data from reference 81

59

An atlas of human blastocysts 60

Figure 2.5 Non-invasive assessment of blastocyst nutrient uptake and utilization. Blastocysts are incubated individually in a known volume, e.g. 0.5 µl, of defined medium such as G277. Serial nanoliter samples can then be taken and analyzed for carbohydrates82, amino acids83, ammonium84, oxygen22 and enzymes85. The concomitant measurement of glucose consumption and lactate production can give an indirect measure of glycolytic activity. Glycolytic activity has been shown to be inversely related to both development in culture and subsequent viability. The concomitant measurement of amino acid consumption and ammonium production can give an indirect measure of amino acid utilization. The release of enzymes, such as lactate dehydrogenase, into the surrounding culture medium reflects impairment in membrane integrity, and as such may be useful in assessing freezing damage

Metabolic requirements during preimplantation development

Figure 2.6 Coupling of enzymatic reaction involving the pyridine nucleotides NAD(P)H (reduced nicotinamide-adenine dinucleotide (phosphate)). Under the appropriate conditions, reactions favor the production of either NADH or NADPH, both of which fluoresce when excited by light in the UV range. When these nucleotides are oxidized (i.e. NAD+ or NADP+), they do not fluoresce. Therefore, in the presence of increasing concentrations of both glucose and lactate, there is a concomitant increase in fluorescence

61

An atlas of human blastocysts 62

generated by the reaction. The absolute amount of fluorescence can then be calibrated daily by running standards and the concentration of substrates present in the culture medium surrounding the embryo determined. For a more detailed account of substrate assays and the reaction conditions required, see reference 82

Figure 2.7 Box plots of glucose uptake (a) and lactate production (b) by individual bovine blastocysts after thawing. Blastocysts were classified retrospectively as either ‘viable’ or ‘non-viable’ based upon their ability, or otherwise, to re-expand within 14 h post-thaw. The lines across the boxes

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represent the median, notches representing the interquartile range, therefore including 50% of the data. Whiskers represent 5 and 95% quartiles. After reference 89

Figure 2.8 Fetal development of mouse blastocysts selected prospectively for transfer using glycolytic activity as a metabolic marker of development. ‘Viable’ blastocysts were those classified with a glycolytic activity close to that of in vivo-developed blastocysts, while ‘non-viable’ blastocysts were those with a highly elevated glycolytic rate. On each day of experiment, a selection of blastocysts were transferred at random. a, b, c: different letters indicate significantly different populations (p<0.01). Data from reference 90

63

An atlas of human blastocysts 64

Figure 2.9 Nutrient uptake and ammonium production by individual human blastocysts from the same patients. Data are in the form of a notched box plot where the notches represent the interquartile range, therefore including 50% of the data points. Whiskers represent 5 and 95% quartiles. The line across the box represents the median. Pyruvate (solid box), glucose (open box), ammonium (hatched box). After reference 94

Figure 2.10 Photomicrograph of a human blastocyst cultured in

Metabolic requirements during preimplantation development

sequential media. Note the development of the ICM and formation of a cohesive epithelium by the trophectoderm cells. Such a blastocyst has an implantation potential (fetal heart rate) of 70%100. The scoring of human blastocysts is strongly related to implantation and pregnancy outcome100,101

Figure 2.11 Schematic diagram of an embryo perfusion culture system. Culture media are continuously passed over the embryo(s). The composition of the culture media can be changed according to the specific requirements of each stage of embryonic development. Toxins such as ammonium are not able to build up and impair embryo development, while more labile components of the culture system are not denatured. Modified from reference 102

65

3 Human morulae in vitro The mammalian morula The word morula derives from the root morus, the Latin word for ‘mulberry’. It was so named in the early days of embryological investigation because of its berry-like shape and appearance, especially notable in amphibians1. The human preembryo is said to have become a morula at approximately the 8-cell stage of development, when the process of compaction commences2. In the human, up to 16 discrete blastomeres may be observed in growing preembryos, but the existence of such high cell numbers without evidence of compaction is now recognized as detrimental to further development (Figure 3.1). Normal compaction results in the formation of an outer layer of cells that become the trophectoderm of the blastocyst, while inner cells give rise to the inner cell mass (insideoutside theory) (Figure 3.2). The embryonic genome becomes functionally active during, or just before, the morula stage. Compaction Compaction is a process that involves the formation of intercellular tight junctions between blastomeres, which become closely apposed and flatten out as the areas of cell contact increase (Figure 3.3). As a result, a cleaving preembryo changes then from a collection of individual cells into a relatively smooth mass with indistinguishable cell outlines (Figure 3.4), the surface of the compacting human morula being characterized by densely distributed microvilli3. Contacts and junctions between mammalian blastomeres are dynamic, and change to accommodate the loss of coupling during mitosis4. It appears that such coupling is mandatory for further development, as demonstrated by the observation that inhibited compaction in mice is lethal for the morula5. The first evidence of compaction is reflected in a polarization of peripheral blastomeres and a reorganization of cytoplasmic components2. Blastomeres or fragments that are unable to form contacts or to communicate appropriately with other blastomeres are generally excluded from the compacting preembryo, often remaining inside the zona pellucida after blastocyst hatching (Figure 3.5). As compaction occurs, cells lose their totipotency as a result of their interactions with one another, and it is believed that this marks the beginning of embryonic DNA transcription. The time of the onset of compaction varies among species: mouse and rat at the 8-cell stage6, bovine and rhesus monkey at the 16–32-cell stage7,8, rabbit at 32–64 cells8, and pig shortly before blastocyst formation9. Human compaction begins after the third mitotic division (8-cell stage) on day 3 and is generally completed by day 4, earlier than in other primate species3. What triggers compaction is unknown, but a developmental ‘clock’ is suggested as being responsible. The process of compaction does not appear to require a

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strict number of cell divisions, and has been observed to begin before completion of the third cleavage in vitro, particularly in slowly developing preembryos (Figure 3.6). In the mouse, inhibition of protein synthesis results in premature compaction10. Similarly, removal of cytoplasm from 1-cell mouse embryos results in accelerated compaction, cavitation and blastocyst formation, possibly by reducing some inhibitory factor11. Conversely, adding cytoplasm from 1-cell zygotes to cleaved embryos slows subsequent development12. These observations suggest that, at least in the mouse, altering nuclear/cytoplasmic ratios by adding or removing cytoplasm may ultimately impact on developmental rates. During compaction, a variety of junctions are formed between cells, in sequential order: gap junctions, adherens junctions, tight junctions and desmosomes (Figures 3.7 and 3.8). Each type plays a fundamental role in cellular communication, adhesion and differentiation13. Gap junctions are composed of a membrane protein named connexin. A group of six connexins form connexson, which in turn forms an intracellular channel: a gap junction is basically a group of intracellular channels14. These transmembrane pores/channels serve as communicators between neighboring cells for transport of metabolites and molecules that regulate cell division, differentiation and apoptosis. Gap junction formation first occurs at the 8-cell stage in the mouse15, while, in the human, the onset and function of these junctions vary from the 4-cell stage to the early blastocyst stage16,17. Electron microscopic studies performed on human morulae show small gap junctions (2–3 nm) within the tight, junctional regions. However, it is unclear whether those of human preblastocyst stages are yet functional, since fluorescent dyes do not pass between cells before the blastocyst stage18. In one of the few studies specific to intracellular junctions in human preembryos, Hardy and colleagues demonstrated expression of the connexin proteins Cx43, Cx32 and Cx26 in 4-cell to hatched blastocyst stages17. Expression of Cx43 was enhanced with increasing development, peaking at the blastocyst stage. Cells of the trophectoderm were noted to be connected by frequent gap junctions, while inner cell masses showed fewer punctate junctions. Abnormal gap junction formation was theorized by these authors to be correlated with reduced conceptus viability. In earlier work involving the microinjection of antibodies against connexin into mouse embryos, the formation of gap junctions was inhibited, causing blastomeres to be extruded19. Such uncoupling has also been observed after induction of low intracellular pH, inhibiting gap junction formation20. Similar factors may be responsible for ‘partial or incomplete’ compaction, frequently observed in buman preembryos cultured in vitro (Figure 3.9). Adherens junctions include the E (epithelial)-cadherin (uvomorulin) system that is calcium-dependent21. In the mouse, E-cadherin is expressed in oocytes and preimplantation embryos during cleavage. Cytoplasmic localization of E-cadherin switches to the membrane regions of adjacent blastomeres in 8-cell specimens, where they mediate early stages of compaction22. At the blastocyst stage, E-cadherin is expressed within trophectoderm and inner cell mass, and operates via the E-cadherin connection to the actin cytoskeleton23. Antibodies against E-cadherin inhibit compaction; however, they do not inhibit gap junction communication24. Furthermore, mouse preimplantation embryos lacking E-cadherin fail to compact, and are rendered non-viable due to an undeveloped trophectoderm25, possibly as a result of disrupted blastomere polarization during the process of trophectoderm differentiation. In the human, E-

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cadherin has been described as being expressed in oocytes and cleavage stage preembryos, but limited to the cell surface26,27. Other investigators demonstrated cytoplasmic expression of E-cadherin in early preimplantation stages, with trophectoderm localization in the blastocyst28. It will be important to investigate possible relationships between failure of compaction and/or blastocyst formation in vitro and disturbance in Ecadherin expression in the human. Tight junctions join adjacent plasma membranes by way of interlinked rows of integral membrane proteins, creating a seal impermeable to the intercellular passage of molecules. They may also result in limited membrane fusion between cells, a situation essential for blastocoel formation (see ‘Cavitation,’ below). Tight junctions interact cytoplasmically with the actin cytoskeleton. The membrane proteins responsible for the tight junctions that form a permanent seal between cells include occludin and claudin; the cytoplasmic proteins include ZO-1, ZO-2 and cingulin29. In addition to forming a seal between cells, tight junctions help to maintain cell polarity, necessary for blastocyst formation and differentiation30. In the mouse, tight junctions begin to develop at the late 8-cell stage after compaction, and require previous establishment of the E-cadherin system. The association between tight junctions and E-cadherin was confirmed by experiments using anti-E-cadherin antibody, where tight junction formation was disrupted29. Tight junction assembly is regulated in stages during the compaction process31. The first stage involves cytoplasmic tight junction proteins, particularly the ZO-1α− variant, which is regulated by E-cadherin and may be responsible for maintaining cell membrane polarity29. In the next stage, cingulin assembles in the apicolateral region of the blastomeres. The final stage of membrane assembly involves the ZO-1α+ variant and the membrane protein, occludin. Formation of ZO-1α+ is thought to be a critical final step in creation of the permeable seal, a prerequisite for cavitation and blastocoel development13,18. Interestingly, delayed expression of ZO-1α+ may prevent occludin transformation, and this delay is possibly involved in developmental arrest of the preembryo. Primitive tight junctions have been observed ultrastructurally in 6–8-cell human preembryos16,32. Other studies have shown that tight junctions are well assembled by the morula stage, and are localized to trophectoderm in the developing human blastocyst18 (Figure 3.10). The expression of ZO-1 proteins in human cleaved preembryos and blastocysts was identified in all stages in the cytoplasm, and localized in trophectoderm junctions in the blastocyst28. Desmosomes are small disk-shaped junctions that connect epithelial cells33. As with tight junctions, key genes regulate the timing of desmosome expression. It appears that desmosomes help to maintain trophectoderm integrity and stability during blastocyst expansion. Premature desmosome development may interfere with differentiation of the trophectoderm and development of the inner cell mass13. In the mouse, desmosomes can be observed between blastomeres and trophectoderm cells34. The same is found in human investigations, where single desmosomes have been identified at the 16-cell stage, with extensive desmosome formation occurring between developing trophectoderm cells during cavitation. In the blastocyst, desmosomes are located between trophectoderm cells but are not found between cells of the inner cell mass or between trophectoderm cells that contact the inner cell mass17 (see Figure 3.8). Ca2+ and Mg2+ ions are also essential for compaction. Compacting human morulae can be decompacted easily by incubation in Ca2+- and Mg2+-free medium for only a few

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minutes (Figure 3.11). This method facilitates blastomere biopsy on day 3, involving 6– 8-cell stage preembryos that have begun to show cell-cell contact and compaction. Other factors that may be involved in the process of compaction include carbohydrate antigens, especially the stage-specific embryonic antigen-1 (SSEA-1). This antigen is expressed on the cell surface of mouse embryos after the third cleavage, and is related to the onset of compaction35. To investigate the possible role of SSEA-1 during the compaction process, specific oligosaccharides were purified from human milk and added to fully compacted mouse morulae. It was found that blastomeres lost cell-cell contacts and became decompacted after 6–8 h36. One possible means of explaining how carbohydrate antigens regulate compaction is to study the Ca2+- and Mg2+-dependent carbohydrate-carbohydrate interaction between cells37. At Cornell, we demonstrated expression of the SSEA-1 antigen on both mature human oocytes and cleaved preembryos up to the blastocyst stage of development. The role of SSEA antigens during human embryogenesis and differential expression between mouse and human are discussed in Chapter 4. The presence of specific adhesion molecules and various junctional complexes in human preembryos indicates the complicated nature of the compaction process. Cavitation After compaction, and as the preembryo expands, it begins to form a cavity. Cavitation is an essential developmental stage whereby cells differentiate into trophectoderm and inner cell mass. As the extent of cell-cell surface contact increases during compaction, the group of cells located on the outside polarize to form the trophectoderm. The smaller group of cells inside develop into the inner cell mass (see Chapter 4). Cavitation involves the accumulation of blastocoelic fluid transported by trophectoderm cells. To accomplish this, trophectoderm cells first depend on complete cellular polarization and then the formation of permeable seals provided by tight junctions which form a belt-like circular line between developing trophectoderm cells38. The polarity of blastomeres is regulated by the extent of their contact and location within the preembryo. Cell polarization involves cell surface microvilli and leptin modifications3,39, irregular allocation of the membrane proteins between apical and basal cell membranes40 and basal mitochondrial accumulation41. The apical membranes of the outer blastomeres of mouse preimplantation embryos possess transport channels that regulate Na+ passage42, while tight junction proteins, gap junctions and E-cadherin are associated with basal membranes31,43,44. In human preembryos, gap junctions are associated with basal and lateral membranes of adjacent blastomeres, while trophectoderm cells of the blastocyst show gap junction formation across their entire surfaces17. The irregular contact between adjacent cells maintains the cell polarity necessary for polarized ion transportation: ions flow in through apical membranes and out through basal membranes into the blastocoel45. Accumulation of fluid into the blastocoel is a result of Na+ transport through the epithelial-like cells of the trophectoderm, and is energy dependent46 (Figure 3.12). Sodium ions diffuse into the trophectoderm through multiple apical channels42. Na/Kadenosine triphosphatase (ATPase), located basolaterally, actively pumps intracellular Na+ into the blastocoel, and tight junctions moderate containment47. Alpha-1 and β-1

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subunits of Na/K-ATPase are expressed during mouse preimplantation development, β subunits being essential for the onset of cavitation48,49. Consequently, blastocyst expansion is inhibited in the presence of ouabain (an inhibitor of Na/K-ATPase)50. Furthermore, ouabain promotes the exchange of intracellular K+ for extracellular Na+, indicating that Na/K-ATPase operates in trophectoderm cells in a manner similar to that in, for example, kidney cells51,52. It is unknown precisely how intracellular Na+ levels are regulated in the trophectoderm, but it is likely that other ions are involved, e.g. Cl−. Apparently, the osmotic gradient resulting from increased Na+ initiates passive water movement into the blastocoel. This is achieved through ‘water channels’ that depend on aquaporins, inherent membrane proteins that allow water to pass in the direction of the osmotic gradient53. Aquaporins are known to be expressed in mouse preimplantation embryos and blastocysts54,55. Intracellular calcium may also play a role in cavitation, since agents such as ethanol and ionomycin that increase intracellular Ca2+ also speed the process of cavitation56,57. Blastocoelic fluid serves as a culture medium during the crucial development of the inner cell mass58,59. Cavitation is clearly an energy-dependent process, and glucose serves as a principal energy source in mammalian preembryos after the 8-cell stage. Cavitation and blastocyst formation in vitro can be greatly influenced by medium composition. Amino acids and other metabolites directly influence normal compaction, cavitation and blastocyst development (see Chapter 2). In the mouse, cavities are formed to one side of the morula, at the so-called abembryonic pole (Figure 3.13). In the human, cavities appear to develop more centrally (Figure 3.14). Morphology of the morula The human morula has generated renewed interest with the routine application of extended culture within clinical in vitro fertilization (IVF) laboratories. Until recently, reports of transferring preembryos on day 4 were limited to cases that involved preimplantation genetic diagnosis (PGD), necessitating a delay of transfer until diagnostic results were obtained. A recent publication contends that this stage of development has been neglected as a transfer option for intrauterine transfer following assisted reproductive technology60. The authors demonstrated increased clinical pregnancy rates following the replacement of compacted morulae, and proposed a grading system for day-4 preembryos. On day 4, compacted morulae were assessed for early compaction (identified by blastomeres that had begun to form a clustered cell mass, each individual cell being identifiable, but not distinct), full compaction (blastomeres were completely adherent, cell boundaries might not be visible, but nuclei could be identified) and late compaction/early blastocyst (cell boundaries were visible again, and cell number was significantly increased). Preembryos not showing any evidence of compaction on day 4 were considered slow-growing and not evaluated in the study. Compacted morulae were further assessed for: (1) The proportion of blastomeres undergoing compaction; (2) The morphology of compaction; (3) Previous morphology on days 2 and 3 of development;

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(4) The percentage of fragmentation. The authors then examined subsequent clinical pregnancy rates based on whether or not high-scoring morulae were replaced. When no high-scoring morulae were available, clinical pregnancy was 28.9% and implantation was 15.7%. Ongoing pregnancy was 22.2%. When at least one ‘good’ morula was transferred, pregnancy, ongoing pregnancy and implantation rates were 50.9%, 37.5% and 25.2%, respectively. When two or more ‘good’ morulae were transferred, rates were 70.0%, 44.9% and 44.4%, respectively. From these data, the authors concluded that day-4 morulae represent a developmental stage with good selection value for intrauterine replacement. Under the light microscope, imminent compaction is evidenced by increased cell-cell contact between blastomeres. Individual blastomeres become closely apposed but still remain separate (Figures 3.15–3.17). As the conceptus begins actively compacting, individual blastomeres become extremely difficult to distinguish as cells begin to flatten and become closely adherent to one another (Figures 3.18–3.20). The fully compacted morula may appear as a single unit exhibiting multiple nuclei. Sometimes individual blastomeres are excluded from the process; often, cytoplasmic fragments are left behind (Figures 3.21–3.23). Early cavitation is observed when a sickle-cell-shaped hollowed area or cleft begins to form between fullyadherent blastomeres. As the conceptus continues its development, a true cavity is formed and begins enlarging. For definition purposes, a cavitating morula is said to possess a cavity constituting less than 50% of its total surface area (Figures 3.24– 3.26). Morulae have been seen to vacuolate (rather than cavitate) in vitro, roughly approximating the appearance of a poorly developing blastocyst, but these soon arrest in their development, secrete no human chorionic gonadotropin (hCG) and have fewer nuclei61 (Figures 3.27–3.29). Human morulae can be observed as early as 65 h post-insemination, but are generally noted some hours after that, between days 3 and 4 of development.

References 1. O’Rahilly R, Mèuller F, Streeter GL. Developmental stages in Human Embryos: Including a Revision of Streeter’s ‘Horizons’ and a Survey of the Carnegie Collection. Washington, DC: Carnegie Institution of Washington, 1987 2. Hartshorne GM, Edwards RG. Early embryo development. In Adashi EY, Rock JA, Rosenwaks Z, eds. Reproductive Endocrinology, Surgery, and Technology. Philadelphia: Lippincott-Raven, 1996:435–50 3. Nikas G, Ao A, Winston RM, Handyside AH. Compaction and surface polarity in the human embryo in vitro. Biol Reprod 1996; 55:32–7 4. Goodall H, Maro B. Major loss of junctional coupling during mitosis in early mouse embryos. J Cell Biol 1986; 102:568–75 5. Cheng SS, Costantini F. Morula decompaction (mdn), a preimplantation recessive lethal defect in a transgenic mouse line. Dev Biol 1993; 156:265–77 6. Reeve WJ. Cytoplasmic polarity develops at compaction in rat and mouse embryos. J Embryol Exp Morphol 1981; 62:351–67

An atlas of human blastocysts 72 7. Enders AC, Lantz KC, Schlafke S. The morula-blastocyst transition in two Old World primates: the baboon and rhesus monkey. J Med Primatol 1990; 19:725–47 8. Koyama H, Suzuki H, Yang X, Jiang S, Foote RH. Analysis of polarity of bovine and rabbit embryos by scanning electron microscopy. Biol Reprod 1994; 50:163–70 9. Reima I, Lehtonen E, Virtanen I, Flechon JE. The cytoskeleton and associated proteins during cieavage, compaction and blastocyst differentiation in the pig. Differentiation 1993;54:35–45 10. Levy JB, Johnson MH, Goodall H, Maro B. The timing of compaction: control of a major developmental transition in mouse early embryogenesis. J Embryol Exp Morphol 1986; 95:213– 37 11. Feng YL, Gordon JW. Removal of cytoplasm from one-celled mouse embryos induces early blastocyst formation. J Exp Zool 1997; 277:345–52 12. Lee DR, Lee JE, Yoon HS, Roh SI, Kim MK. Compaction in preimplantation mouse embryos is regulated by a cytoplasmic regulatory factor that alters between 1- and 2-cell stages in a concentration-dependent manner. J Exp Zool 2001; 290:61–71 13. Fleming TP, Ghassemifar MR, Sheth B. Junctional complexes in the early mammalian embryo. Semin Reprod Med 2000; 18:185–93 14. Kidder GM, Winterhager E. Intercellular communication in preimplantation development: the role of gap junctions. Front Biosci 2001; 6:D731–6 15. Lo CW, Gilula NB. Gap junctional communication in the preimplantation mouse embryo. Cell 1979; 18:399–409 16. Dale B, Gualtieri R, Talevi R, Tosti E, Santella L, Elder K. Intercellular communication in the early human embryo. Mol Reprod Dev 1991 ;29:22–8 17. Hardy K, Warner A, Winston RM, Becker DL. Expression of intercellular junctions during preimplantation development of the human embryo. Mol Hum Reprod 1996; 2:621–32 18. Gualtieri R, Santella L, Dale B. Tight junctions and cavitation in the human preembryo. Mol Reprod Dev 1992; 32:81–7 19. Lee S, Gilula NB, Warner AE. Gap junctional communication and compaction during preimplantation stages of mouse development. Cell 1987; 51:851–60 20. Leclerc C, Becker D, Buehr M, Warner A. Low intracellular pH is involved in the early embryonic death of DDK mouse eggs fertilized by alien sperm. Dev Dyn 1994; 200:257–67 21. Takeichi M. Cadherin cell adhesion receptors as a morphogenetic regulator. Science 1991; 251:1451–5 22. Clayton L, Stinchcombe SV, Johnson MH. Cell surface localisation and stability of uvomorulin during early mouse development. Zygote 1993; 1:333–44 23. Aberle H, Schwartz H, Kemler R. Cadherin-catenin complex: protein interactions and their implications for cadherin function. J Cell Biochem 1996; 61:514–23 24. Johnson MH, Maro B, Takeichi M. The role of cell adhesion in the synchronization and orientation of polarization in 8-cell mouse blastomeres. J Embryol Exp Morphol 1986; 93:239– 55 25. Riethmacher D, Brinkmann V, Birchmeier C. A targeted mutation in the mouse E-cadherin gene results in defective preimplantation development. Proc Natl Acad Sci USA 1995; 92:855–9 26. Campbell S, Swann HR, Seif MW, Kimber SJ, Aplin JD. Cell adhesion molecules on the oocyte and preimplantation human embryo. Hum Reprod 1995; 10:1571–8 27. Rufas O, Fisch B, Ziv S, Shalgi R. Expression of cadherin adhesion molecules on human gametes. Mol Hum Reprod 2000; 6:163–9 28. Bloor DJ, Metcalfe AD, Rutherford A, Brison DR, Kimber SJ. Expression of cell adhesion molecules during human preimplantation embryo development. Mol Hum Reprod 2002; 8:237– 45 29. Fleming TP, McConnell J, Johnson MH, Stevenson BR. Development of tight junctions de novo in the mouse early embryo: control of assembly of the tight junction-specific protein, ZO1. J Cell Biol 1989; 108:1407–18

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30. Stevenson BR, Keon BH. The tight junction: morphology to molecules. Annu Rev Cell Dev Biol 1998; 14:89–109 31. Fleming TP, Sheth B, Fesenko I. Cell adhesion in the preimplantation mammalian embryo and its role in trophectoderm differentiation and blastocyst morphogenesis. Front Biosci 2001; 6:D1000–7 32. Tesarik J. Involvement of oocyte-coded message in cell differentiation control of early human embryos. Development 1989; 105:317–22 33. Garrod D, Chidgey M, North A. Desmosomes: differentiation, development, dynamics and disease. Curr Opin Cell Biol 1996; 8:670–8 34. Fleming TP, Garrod DR, Elsmore AJ. Desmosome biogenesis in the mouse preimplantation embryo. Development 1991; 112:527–39 35. Eggens I, Fenderson B, Toyokuni T, Dean B, Stroud M, Hakomori S. Specific interaction between Lex and Lex determinants. A possible basis for cell recognition in preimplantation embryos and in embryonal carcinoma cells. J Biol Chem 1989; 264:9476–84 36. Fenderson BA, Eddy EM, Hakomori S. Glycoconjugate expression during embryogenesis and its biological significance. Bioessays 1990; 12:173–9 37. Solter D, Knowles BB. Monoclonal antibody defining a stage-specific mouse embryonic antigen (SSEA-1). Proc Natl Acad Sci USA 1978; 75:5565–9 38. Watson AJ, Barcroft LC. Regulation of blastocyst formation. Front Biosci 2001; 6:D708–30 39. Antczak M, Van Blerkom J. Oocyte influences on early development: the regulatory proteins leptin and STAT3 are polarized in mouse and human oocytes and differentially distributed within the cells of the preimplantation stage embryo. Mol Hum Reprod 1997; 3:1067–86 40. Watson AJ. The cell biology of blastocyst development. Mol Reprod Dev 1992; 33:492–504 41. Wiley LM. Cavitation in the mouse preimplantation embryo: Na/K-ATPase and the origin of nascent blastocoele fluid. Dev Biol 1984; 105:330–42 42. Manejwala FM, Cragoe EJ Jr, Schultz RM. Blastocoel expansion in the preimplantation mouse embryo: role of extracellular sodium and chloride and possible apical routes of their entry. Dev Biol 1989; 133:210–20 43. Kemler R. From cadherins to catenins: cytoplasmic protein interactions and regulation of cell adhesion. Trends Genet 1993; 9:317–21 44. Vestweber D, Gossler A, Boller K, Kemler R. Expression and distribution of cell adhesion molecule uvomorulin in mouse preimplantation embryos. Dev Biol 1987; 124:451–6 45. Nuccitelli R, Wiley LM. Polarity of isolated blastomeres from mouse morulae: detection of transcellular ion currents. Dev Biol 1985; 109:452–63 46. Biggers JD, Bell JE, Benos DJ. Mammalian blastocyst: transport functions in a developing epithelium. Am J Physiol 1988; 255:C419–32 47. Watson AJ, Kidder GM. Immunofluorescence assessment of the timing of appearance and cellular distribution of Na/K-ATPase during mouse embryogenesis. Dev Biol 1988:126:80–90 48. Watson AJ, Pape C, Emanuel JR, Levenson R, Kidder GM. Expression of Na,K-ATPase α and β subunit genes during preimplantation development of the mouse. Dev Genet 1990:11:41–8 49. Gardiner CS, Williams JS, Menino AR Jr. Sodium/potassium adenosine triphosphatase a- and β-subunit and a-subunit mRNA levels during mouse embryo development in vitro. Biol Reprod 1990; 43:788–94 50. DiZio SM, Tasca RJ. Sodium-dependent amino acid transport in preimplantation mouse embryos. III. Na+-K+-ATPase-linked mechanism in blastocysts. Dev Biol 1977; 59:198–205 51. Baltz JM, Smith SS, Biggers JD, Lechene C. Intracellular ion concentrations and their maintenance by Na+/K+-ATPase in preimplantation mouse embryos. Zygote 1997; 5:1–9 52. Watson AJ, Damsky CH, Kidder GM. Differentiation of an epithelium: factors affecting the polarized distribution of Na+,K+-ATPase in mouse trophectoderm. Dev Biol 1990; 141:104–14 53. Deen PM, van Os CH. Epithelial aquaporins. Curr Opin Cell Biol 1998; 10:435–42 54. Edashige K, Sakamoto M, Kasai M. Expression of mRNAs of the aquaporin family in mouse oocytes and embryos. Cryobiology 2000; 40:171–5

An atlas of human blastocysts 74 55. Offenberg H, Barcroft LC, Caveney A, Viuff D, Thomsen PD, Watson AJ. mRNAs encoding aquaporins are present during murine preimplantation development. Mol Reprod Dev 2000; 57:323–30 56. Stachecki JJ, Armant DR. Regulation of blastocoele formation by intracellular calcium release is mediated through a phospholipase C-dependent pathway in mice. Biol Reprod 1996; 55:1292–8 57. Stachecki JJ, Armant DR. Transient release of calcium from inositol 1,4,5-trisphosphatespecific stores regulates mouse preimplantation development. Development 1996; 122:2485–96 58. Dardik A, Schultz RM. Protein secretion by the mouse blastocyst: differences in the polypeptide composition secreted into the blastocoel and medium. Biol Reprod 1991; 45:328–33 59. Dardik A, Doherty AS, Schultz RM. Protein secretion by the mouse blastocyst: stimulatory effect on secretion into the blastocoel by transforming growth factor-α. Mol Reprod Dev 1993; 34:396–401 60. Tao J, Tamis R, Fink K. Pregnancies achieved after transferring frozen morula/compact stage embryos. Fertil Steril 2001; 75:629–31 61. Dokras A, Sargent IL, Barlow DH. Human blastocyst grading: an indicator of developmental potential? Hum Reprod 1993; 8:2119–27

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Figure 3.1 Day-3 preembryos that fail to begin compaction by the 8-cell stage of development. (a) Human preembryo on day 3 with 16 blastomeres, displaying no evidence of compaction. Neither compaction nor blastocyst development occurred over the following 3 days; (b) day-3 human preembryo with 14–16 blastomeres, shown here immediately before

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transfer, that failed to implant after intrauterine transfer

Figure 3.2 Differentiation of inner cell mass (ICM) and trophectoderm (TM) following compaction. (a) Separation of cell types and early cavitation; central inner cell mass and peripheral trophectoderm. The blastocoele can be seen forming between the ICM and TM, occupying nearly 50% of the surface area; (b) morula at more advanced stage of cavitation. Clear formation of the ICM, its migration from a central to more polar position,

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and a peripheral TM. ICM cells are rounded while TM cells are elliptical and polarized. Blastocoele encompasses approximately 50% of the surface area; (c) centrallydeveloping ICM shows dispersed cells beginning to join together; (d) fully expanded blastocyst with compacted ICM and cohesive, multicellular TM

Figure 3.3 Human preembryos during initial stages of compaction, termed the stage of cell-cell contact. Blastomeres become closely apposed to one another and cell junctions begin to form

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between them. (a)–(d) Early cell-cell contact when blastomeres may still be counted easily

Figure 3.4 Human preembryos exhibiting increased cell-cell contact. As the areas of cell contact increase, individual cell membranes are less defined. (a) Blastomeres are closely apposed but can still be counted; (b) with increased contact, blastomeres cannot be counted easily; (c) and (d) fully compacted morula; cell membranes cannot be distinguished but nuclei can be counted

Figure 3.5 Blastomeres and fragments unable to form appropriate contacts are generally excluded from the compaction process. (a) Compacted morula at the onset of cavitation. Fragments and/or blastomeres not participating in the process are shown to the right; (b) two to three large, vacuolating blastomeres to the left are excluded from compaction and

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cavitation; (c) and (d) cells and/or fragments excluded from the blastocyst remain in the zona pellucida after hatching

Figure 3.6 Compaction in slowly developing preembryos before the 8cell stage. (a)–(e) The fact that compaction occurs in slowly

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developing human preembryos (4–7cell stages) on day 4 supports the idea of a developmental clock regulating the process. These preembryos usually do not form viable blastocysts

Figure 3.7 Schematic presentation of cell junctions. (a) Gap junctions are comprised of connexins that form gaps

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between blastomere membranes. Adherens junctions involve the cadherin system connected to actin filaments via catenins; (b) tight junctions are extremely close connections formed between membranes of TM cells. Desmosomes are disk-shaped junctions formed between TM cells

Figure 3.8 Transmission electron micrograph of human blastocyst showing cell junctions between two trophectoderm cells, i.e. tight junction and desmosome (arrows). Photograph courtesy of Henry A.Sathananthan

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Figure 3.9 Partial/incomplete compaction. (a) Partial compaction frequently seen in human preembryos cultured in vitro. One or more blastomeres of the preembryo will not participate in the compaction process; (b) exclusion of one large, vacuolated blastomere does not impede compaction of the remainder of the conceptus

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Figure 3.10 High-power magnification of the trophectoderm cells of a human blastocyst. TM cells are elongated, elliptical in shape, and have polarized membranes. Tight junctions mediate connections between these cells

An atlas of human blastocysts 84

Figure 3.11 Human morula exposed briefly to Ca2+- and Mg2+-free medium. (a) Compacted morula exhibiting some excluded cells before treatment; (b) morula after treatment showing decompaction as evidenced by the loss of blastomere coupling

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Figure 3.12 Schematic representation of the cavitation process. TM cells are polarized apically, facing the zona pellucida; basal membranes face the blastocoele. Tight junctions between TM cells form permeable seals necessary for fluid accumulation. Sodium ions diffuse through TM cells via sodium channels. NA/K-ATPase, located basolaterally, is responsible for the active pumping of intracellular sodium into the blastocoele. The resulting osmotic gradient initiates passive water movement into the blastocoele. Tight junctions moderate containment

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Figure 3.13 Exocentric cavitation of the mouse morula. The cavity forms at the abembryonic (AB) pole, while the ICM forms at the embryonic (EB) pole. The AB/EB orientation depends upon polar body (PB) position

Figure 3.14 Central cavitation of the human morula. The cavity forms centrally, surrounding ICM cells as they separate from the developing TM. There are notable differences between

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mouse (Figure 3.13) and human cavitation

Figure 3.15 (a)–(f) Human preembryos on day 3 (8–10 blastomeres) exhibiting minimal cellcell contact. Blastomeres are counted easily

An atlas of human blastocysts 88

Figure 3.16 (a)–(f) Human preembryos on day 3 showing minimal to moderate cell-cell contact. Membranes of blastomeres can still be seen clearly

Figure 3.17 (a) and (b) Moderate cellcell contact between blastomeres

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Figure 3.18 (a)–(d) Full cell-cell contact between blastomeres (compaction). Individual blastomere membranes are becoming difficult to recognize

Figure 3.19 (a)–(d) Compacting human morulae on day 4. Tightly compacted cells are observed.

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Compacted human morulae on day 4 showing multiple nuclei of different sizes and shapes, indicating possible multinucleation

Figure 3.20 (a) and (b) Human morulae at the final stage of compaction on day 4. Individual cell membranses are no longer visible

Figure 3.21 (a)–(c) Fully compacted human morulae on day 4; first evidence of minor decompaction just before the onset of cavitation. Cell membranes become somewhat visible once again. One may observe either cytoplasmic accumulations around cell nuclei or areas completely clear of cytoplasm (at presumed blastomere peripheries). If the latter areas increase,

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cavitation and subsequent blastocyst development may be impaired. This may indicate intracytoplasmic, instead of extracytoplasmic, water accumulation, possibly stemming from an abnormal early cavitation process (Na/K-ATPase)

Figure 3.22 (a) and (b) Further minor decompaction; extremely small clefts are appearing between cells, indicating the very beginning of cavitation

Figure 3.23 Compaction → decompaction → cavitation. (a) Slight decompaction; small clefts indicating onset of cavitation; total cell number is increasing; (b) two decompacting, early cavitating morulae. Some decompaction is taking place and cell number is increasing just before the onset of cavitation; (c) human morula showing increased cell division (thus,

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an increase in cell number) and minor decompaction and clefting just before cavitation

Figure 3.24 (a)–(f) Beginning of cavitation. Small fluid-filled areas are formed between adjacent cells of the morula. In time, these areas will enlarge to form a single cavity, the blastocoele. Cells differ in size and shape. No clear ICM can yet be discerned

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Figure 3.25 (a)–(f) Advancing cavitation in human morulae. Increased accumulation of blastocyst fluid can be observed. Early morphological differences between the rounded cells of the inner cell mass and elliptical cells of the trophectoderm are apparent. The blastocoele still occupies less than 50% of the blastocyst surface area

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Figure 3.26 (a)–(f) Final stage of cavitation in the human morula (Cornell criteria). The cavity occupies approximately 50% of the morula surface area. Although no increase in overall size has yet occurred (no overall expansion), these conceptuses may be called early blastocysts by others, since cells have clearly separated to inner cell mass and trophectoderm components

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Figure 3.27 Abnormal cavitation of the human morula. (a) Cavitation in a morula made up of few blastomeres. Typically, a low cell number is observed in this type of abnormal morula before compaction. While the mechanisms contributing to cavity formation are active (if only in few cells), the future blastocyst is compromised; (b) exocentric, singlesided cavitation, or vacuolization

Figure 3.28 (a) and (b) Typical vacuolization in human morulae. Single, unhealthy appearing cells are pushed to the side due to accumulating fluid. Some individual cells are associated with the fluid-filled area, but overall morphology is quite poor.

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Specimens such as these always arrest at this stage

Figure 3.29 (a) and (b) Vacuolated morulae on day 5. These are completely filled with fluid, although no ICM or TM has formed. There is, however, a notable increase in overall size and the zona pellucida thins under fluid pressures. These conceptuses always arrest and never hatch

4 Cell allocation and differentiation During blastocyst development, two distinguishable cell types are formed, those of the inner cell mass and those of the trophectoderm. Trophectoderm represents the first truly differentiated cell line to be established in the newly formed conceptus, differences between these cells and those of the inner cell mass becoming apparent as the trophectoderm and inner cell mass separate and migrate to their new positions during cavitation (Figure 4.1). During blastocyst formation, trophectoderm cells become elliptical and exhibit polarization, whereas cells of the inner cell mass remain rounded and morphologically undifferentiated (Figure 4.2). Trophectoderm cells connect to one another over small surface areas, while cells of the inner cell mass become tightly compacted and closely apposed. The position and development of the trophectoderm and inner cell mass are not random, but are dependent on genetic, morphological and developmental mechanisms. Regulated at the start of the fertilization process, or even earlier in some mammals, this involves a distinct polarization. In most mammals, including the human, polarization and formation of the embryonic axes are fundamental to mammalian pre- and postimplantation development1. Mammalian oocytes express a polarized organization in which specific maternal proteins, leptin and STAT3, are expressed at the ‘animal’ (anterior) pole of growing oocytes and on trophectoderm cells of blastocysts2. The second polar body position marks the anterior pole, the posterior pole being opposite, and the polar axis marks the position of the first cleavage plane (Figure 4.3). It was suggested by Piotrowska and Zernicka-Goetz, and later questioned by Davies and Gardner, that the mouse oocyte may be polarized in regard to sperm entry; thus, the site of sperm entry is believed to predict the first cleavage plane, and may also predict embryonic and abembryonic regions of the future blastocyst3,4. The human oocyte, in contrast, is not polarized in this manner, and the observation that fragments from human oocytes (cytoplasts) can form a pronucleus after incubation with spermatozoa provides evidence that there is no specific sperm entry site5,6. During fertilization, polarized nucleoli are apparent within the pronuclei, and mitochondria become aggregated around pronuclei7,8. The first cleavage plane in mouse and human zygotes is established by the position of the second polar body, which serves as a marker of the polar axis from zygote to blastocyst stages (Figures 4.4 and 4.5). The anterior-posterior axes and dorsal-ventral axes determine anterior-posterior and embryonic-abembryonic poles (Figures 4.6 and 4.7). These axes are well established in the mouse, but less clear in the human7. In a lineage-tracing study performed in the mouse, totipotent 2-cell stage blastomeres were reported to contribute equally to inner cell mass and trophectoderm cell lines9. The validity of this conclusion has been questioned subsequently however, since a growing

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body of evidence suggests that polarity plays a major role in murine development10. In the mouse at least, blastomeres of the 2-cell stage can develop into either embryonic or abembryonic components. It has been demonstrated further that one blastomere at the 2cell stage cleaves earlier than the other and preferentially contributes to the embryonic part of the developing blastocyst11. Recall here that the onset of embryonic gene expression in the mouse begins at the 2-cell stage, as opposed to the 4–8-cell stages in humans. Experiments on human preembryos, in which a tracing dye was injected into a single blastomere at the 2–8-cell stage, showed later random distribution of dye between inner cell mass and trophectoderm cells12. Future studies must investigate the fate of single blastomere development within the concept of preembryo polarization, possibly by using cell-permeable dyes, since the intracellular injection technique might have been responsible for delaying cell division and altering developmental outcome13. In the human, blastomeres of the 8-cell stage have been thought to be genetically equal, a prerequisite for preimplantation genetic diagnosis (PGD). However, if developmental differences between blastomeres do exist before the third cleavage, then genetic analyses of individual blastomeres at the 8-cell stage will evaluate only one of the future developmental lineages, either inner cell mass or trophectoderm. This concern is supported by studies using Oct4 and β-human chorionic gonadotropin (β-hCG) (markers of the inner cell mass and trophectoderm, respectively), where neither was expressed uniformly within blastomeres of cleavage stage preembryos14,15. Mouse blastomeres at the first cleavage are clearly totipotent, since the two can each develop into a normal blastocyst when separated16. This totipotency appears to become limited soon thereafter, because isolated 4–8-cell blastomeres are often observed to develop abnormally, primarily as trophectoderm17. In humans, however, it is clear from the experience of PGD that removal of one or two blastomeres at the 8-cell stage does not necessarily compromise subsequent development18. Conversely, removal of one blastomere at the 4-cell stage or earlier impairs further development, and results in a reduction in the total number of cells of the inner cell mass19. Various models for the basis of mammalian cell allocation and differentiation have been proposed over the years. These include: the inside-outside hypothesis, the polarization hypothesis and the cleavage-driven hypothesis20 (Figure 4.8). Tarkowski and Wroblewska proposed the inside-outside hypothesis, wherein the fate of blastomeres is described as being a consequence of their differing locations within the preembryo17. These investigators detailed a situation where inner blastomeres of the 8cell stage preembryo are destined to form the inner cell mass and those outside to form the trophectoderm. In accord with this, isolated blastomeres from 4–8-cell mouse and 8cell human preembryos, no longer totipotent (see Glossary), are capable of developing only into trophectoderm-like vesiculated structures, and subsequently fail to form inner cell masses and normal trophectoderm, probably owing to a reduction in cellular mass21. The polarization hypothesis proposes that, during the fourth cleavage to 16 cells, polarized surfaces and a cytoskeleton develop in an outside to inside direction22. Subsequent cleavage generates daughter cells with different developmental fates. The newer cleavage-driven hypothesis includes elements of the previous two hypotheses and incorporates the idea of blastomeres becoming predisposed at the first

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cleavage to contribute to embryonic portions of the blastocyst20. According to this theory, blastocyst polarity arises from early asymmetric cleavage of the preembryo. Each of these hypotheses will require scientific verification in a human model, since the only analyses to date relate to the morphology of early development7. From such investigations carried out in our laboratory, it appears that the human preembryo enters compaction at the 8-cell stage and that further cleavage is inhibited until the compaction process is completed (Figures 4.9 and 4.10). Following compaction, some blastomeres are immediately allocated to the outside, where they begin to proliferate rapidly to form the trophectoderm. This separation of cells is clearly evident during cavitation and subsequent blastocyst formation (Figures 4.11 and 4.12). A few cells remaining inside the morula ultimately form the inner cell mass. Connections between blastomeres, their polarization, and cellular orientation within the preembryo regulate embryonic differentiation (see Chapter 3). The ratio of cells contributing to each cell type is tightly controlled. The blastocyst is polarized, with the inner cell mass localized at the embryonic pole7. The inner cell mass represents a group of cells that are tightly connected and pluripotent (see Glossary), while trophectoderm cells have smaller areas of their surfaces connected, serve as protectors of the inner cell mass and are differentiated for this function. If trophectoderm cells are removed, the morphology of the inner cell mass changes rapidly and pluripotency is lost (Figure 4.13). In the mouse, the inner cell mass regulates trophectoderm development and its distribution within the blastocyst. Fibroblast growth factor-4 has been shown to be involved in this process23. It is unclear just how cells of the inner cell mass direct trophectoderm differentiation and distribution, since daughter cells on the outside of the morula usually form the trophectoderm24. Interestingly, parthenogenic mouse conceptuses develop into blastocysts with lower total cell numbers and lower inner cell mass/trophectoderm cell ratios, implicating gene imprinting as an important factor even during preimplantation development25.

Blastocyst cell number For successful development and implantation, the total number of cells in the blastocyst and the inner cell mass/trophectoderm cell ratio are genetically and developmentally regulated26. The average cell cycle in mouse trophectoderm requires 17.5 h, while that of the inner cell mass is 24 h. Thus, the trophectoderm proliferates more rapidly as compared to the inner cell mass, averaging one extra full cell division per 24 h24,27. In the human also, the trophectoderm proliferates more rapidly as compared to the inner cell mass. Trophectoderm cell numbers double from the onset of cavitation on day 4 to the formation of the blastocyst on day 5, while the numbers in the inner cell mass double between days 5 and 6. During blastocyst expansion, cells of the trophectoderm divide at a rate of one-half a division per 24 h more rapidly than cells of the inner cell mass28. Still, quantitative studies of the human blastocyst demonstrate great variability in total cell number. Generally, total cell numbers in normal human blastocysts should exceed 60 cells. The human blastocyst has approximately 60 cells on day 5, up to 160 cells on day 6 and over 200 cells after hatching on day 728–30. In the mouse, the inner cell mass/trophectoderm ratio changes during blastocyst development from 40:60 to

An atlas of human blastocysts 100

25:75; in the human, approximately 40% of the final cell number is made up of inner mass cells28. The exact mechanisms that control cell number in the human and primate blastocyst are unknown. These mechanisms are conserved during early cleavage stages, as evidenced by the fact that rhesus monkey blastocysts developing after separation of the blastomeres (after artificial twinning) contain half the number of total cells while maintaining the same inner cell mass/trophectoderm ratios31 (see Chapter 12). Apoptotic processes may be one of the possible systems involved in regulating blastocyst cell number32. Quantitative analysis of the total blastocyst cell number can be performed by using various techniques that visualize individual cell nuclei. As an example, the total cell number in living specimens can be evaluated easily using cell permeable fluorescence nuclear dyes, e.g. Hoechst (Figure 4.14). Other methods include fixation and staining with nuclear dyes (propidium iodide, Giemsa® or 4’,6-diamidino-2-phenylindole). To quantify and compare the numbers of cells in the inner cell mass and trophectoderm, differential staining techniques were developed33. Techniques include differential staining of whole specimens, or isolation and staining of the inner cell mass alone34. These techniques are based on the high permeability of the trophectoderm and the relatively poor permeability of the tightly connected inner cell mass. Immunological techniques include the use of a species-specific antiserum followed by complement, with propidium iodide used to stain the nuclei of lysed trophectoderm cells; cells of the inner cell mass are then stained with permeable dyes27. A modification of this technique involves using trinitrobenzenesulfonic acid (TNMS) instead of antiserum28. Differential staining through chemical permeabilization techniques can be achieved using Ca2+ ionophore A23187 in combination with nuclear fluorochromes; however, lysing of trophectoderm cells is a necessary preliminary, based on an osmotic response that causes membrane vesiculation35. Modifications of this technique include the labeling of trophectoderm with fluorescent-labeled lectins, e.g. wheat-germ agglutinin36, or using the detergent Triton X-100® to permeabilize trophectoderm cells37. Blastocyst quality can be evaluated by both non-invasive and invasive techniques. Non-invasive techniques include evaluating the timing of blastocyst expansion, trophectoderm morphology and inner cell mass characteristics. Other non-invasive techniques include the metabolic evaluation of nutrient uptake and/or production. Recently, it was shown that depletion of leucine and formation of alanine are positively correlated with normal blastocyst formation and implantation in the human38. Invasive techniques generally result in a loss of viability. They include, among others, karyotyping, fluorescence in situ hybridization (FISH) for analysis of chromosome structure or number (aneuploidy, mosaicism) and analysis of mitotic or apoptotic indices. Gene expression and regulation The genes involved in blastocyst formation are largely unknown, but are probably induced or regulated by activation of some preembryonic genes at the 4–8-cell stage in the human. Known genes include those for sodium/potassium adenosine triphosphatase and ZO-1α+39,40 (see Chapter 3). The preimplantation embryo development gene (Ped gene) is located in the Q region of the major histocompatibility complex in mice, and has fast- and slow-cleaving forms.

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In more rapidly growing preembryos, this gene induces faster cleavage and a higher total cell number in the subsequent blastocyst41. It has been suggested that human leukocyte antigen-G (HLA-G), expressed in human blastocysts, is homologous to the Ped gene in mice42. As might be expected, gene expression differs between the inner cell mass and the trophectoderm. These differences probably reflect the loss of totipotency and formation of differentiated trophectoderm cells. One well-known marker of totipotency is Oct4. This transcription factor is crucial for maintaining totipotency in early cleaving preembryos, is responsible for maintaining pluripotency in the inner cell mass and is later restricted to germ cells during embryogenesis43. In the absence of Oct4, early lethality occurs due to the differentiation as trophectoderm of those cells that would normally give rise to the inner cell mass44. Another important role for Oct4 is the maintenance of embryonic stem cells in an undifferentiated state, it being down-regulated before cells are triggered to differentiate45. Oct4 is differentially expressed in various species including the human46. In humans, Oct4 is expressed on the oocyte and throughout development to the blastocyst stage47,48. The distribution of Oct4 varies between blastomeres of human preembryos, suggesting a possible directive mechanism towards inner cell mass (Oct4-positive) or trophectoderm (Oct4-negative) lineages14. In the human blastocyst, Oct4 is expressed primarily in the inner cell mass, where mRNA expression is approximately 31 times higher than in the trophectoderm49 (Figure 4.15). Elevated Oct4 expression is maintained in undifferentiated embryonic stem cells as they develop from the inner cell mass. Recently, poor results with somatic cell cloning were associated with abnormal and/or failed expression of Oct4 in cloned mouse embryos and blastocysts50. Trophectoderm-associated genes regulate membrane polarity during the compaction and cavitation stages necessary for normal trophectoderm formation (see Chapter 3). One might be the helix-loop-helix transcription factor (Hxt), a gene that is increasingly expressed in mouse morulae, trophectoderm and placenta51. In addition, transforming growth factor type β2 (TGF-β 2) is expressed in outer cells of the morula and in trophectoderm, while the inner cell mass is unstained52. Also, the β subunit of human chorionic gonadotropin (β-hCG), a trophectoderm marker, is expressed in human cleavage-stage preembryos, and hCG is later secreted from the trophectoderm cells of the blastocyst15,53,54. Carbohydrates are further elements that play a role in preimplantation development. Glycans, which are glycoprotein-rich carbohydrates, are characteristic of early embryonic cells that carry a number of carbohydrate markers. As in other cell systems, surface carbohydrates play a critical role in the cell-cell interactions of mammalian preembryos55,56. Stage-specific embryonic antigens (SSEAs) are surface glycoconjugates associated with glycolipids, glycoproteins and proteoglycans, and are developmentally regulated57. In the mouse, SSEA-3 and SSEA-4 are glycolipid antigens evident during oogenesis, which are expressed in oocytes and cleavage stage preembryos but which disappear by the blastocyst stage58. In contrast, SSEA-1 is expressed from the 8-cell stage, participates in the process of compaction59, is later expressed mainly in the inner cell mass, and in embryonic stem cells serves as a marker of non-differentiation60,61.

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In the human, SSEA-3 and -4 antigens are markers of undifferentiated stem cells, while the presence of SSEA-1 indicates a differentiated state.These species-specific differences have been confirmed by our own studies with human and mouse oocytes and preembryos (Figures 4.16–4.18)62. SSEA-3 and -4 expression becomes restricted to the inner blastomeres of the compacting human morula (support for the inside-outside theory), indicating that human cell differentiation begins by this stage and continues throughout the process of cell allocation. Subsequently, the inner cell mass expresses SSEA-3 and -4, while cells of the trophectoderm do not. Mature human oocytes express SSEA-1 while immature oocytes appear not to; preembryos and blastocysts

Table 4.1 Expression of stage-specific embryonic antigen SSEA: mouse and human SSEA-1

SSEA-3

SSEA-4

Mouse oocyte

−

+

+

Human oocyte

+mature/−immature

+

+

Mouse embryo

−<8-cell/+≥8-cell

+

+

Human preembryo

+

+

+

Mouse blastocyst

+

−

−

inner cell mass

+

−

−

trophectoderm

+

−

−

Human blastocyst

+

+/−

+/−

inner cell mass

+

+

+

trophectoderm

+

−

−

Mouse embryonic stem cell

+

−

−

Human embryonic stem cell

−

+

+

express it consistently. A comparison of SSEA expression in both human and mouse is presented in Table 4.1. Henderson and colleagues recently confirmed most of The differences that they found before the 8-cell stage our results of SSEA expression in human blastocysts63. may be technical in nature. We stained live specimens, as did Solter and Knowles in their original experiments60, while Henderson used fixed material in which we had earlier noted only faint or no staining. The techniques involving SSEA staining in living specimens will allow selection of SSEA-3- and -4-positive inner cell masses for future embryonic stem cell culture, and undifferentiated stem cells for manipulation.

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46. Kirchhof N, Carnwath JW, Lemme E, Anastassiadis K, Scholer H, Niemann H. Expression pattern of Oct-4 in preimplantation embryos of different species. Biol Reprod 2000; 63:1698– 705 47. Abdel-Rahman B, Fiddler M, Rappolee D, Pergament E. Expression of transcription regulating genes in human preimplantation embryos. Hum Reprod 1995; 10:2787–92 48. Verlinsky Y, Morozov G, Verlinsky O, et al. Isolation of cDNA libraries from individual human preimplantation embryos. Mol Hum Reprod 1998; 4:571–5 49. Hansis C, Grifo JA, Krey LC. Oct-4 expression in inner cell mass and trophectoderm of human blastocysts. Mol Hum Reprod 2000;6:999–1004 50. Boiani M, Eckardt S, Scholer HR, McLaughlin KJ. Oct4 distribution and level in mouse clones: consequences for pluripotency. Genes Dev 2002; 16:1209–19 51. Cross JC, Flannery ML, Blanar MA, et al. Hxt encodes a basic helix-loop-helix transcription factor that regulates trophoblast cell development. Development 1995; 121:2513–23 52. Slager HG, Lawson KA, van den Eijnden-van Raaij AJ, de Laat SW, Mummery CL. Differential localization of TGF-(32 in mouse preimplantation and early postimplantation development. Dev Biol 1991; 145:205–18 53. Bonduelle ML, Dodd R, Liebaers I, Van Steirteghem A, Williamson R, Akhurst R. Chorionic gonadotrophin-β mRNA, a trophoblast marker, is expressed in human 8-cell embryos derived from tripronucleate zygotes. Hum Reprod 1988; 3:909–14 54. Fishel SB, Edwards RG, Evans CJ. Human chorionic gonadotropin secreted by preimplantation embryos cultured in vitro. Science 1984; 223:816–18 55. Poirier F, Kimber S. Cell surface carbohydrates and lectins in early development. Mol Hum Reprod 1997; 3:907–18 56. Muramatsu T. Developmentally regulated expression of cell surface carbohydrates during mouse embryogenesis. J Cell Biochem 1988; 36:1–14 57. Fenderson BA, Eddy EM, Hakomori S. Glycoconjugate expression during embryogenesis and its biological significance. Bioessays 1990; 12:173–9 58. Shevinsky LH, Knowles BB, Damjanov I, Solter D. Monoclonal antibody to murine embryos defines a stage-specific embryonic antigen expressed on mouse embryos and human teratocarcinoma cells. Cell 1982; 30:697–705 59. Fenderson BA, Zehavi U, Hakomori S. A multivalent lacto-N-fucopentaose lll-lysyllysine conjugate decompacts preimplantation mouse embryos, while the free oligosaccharide is ineffective. J Exp Med 1984; 160:1591–6 60. Solter D, Knowles BB. Monoclonal antibody defining a stage-specific mouse embryonic antigen (SSEA-1). Proc Natl Acad Sci USA 1978; 75:5565–9 61. Smith AG, Heath JK, Donaldson DD, et al. Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature (London) 1988; 336:688–90 62. Zaninovic N, Veeck LL, Rosenwaks Z. Stage-specific expression of embryonic antigens SSEA1, SSEA-3, and SSEA-4 on human conceptuses using vital staining. Fertil Steril 2001; 76:S33– S34 63. Henderson JK, Draper JS, Baillie HS, et al. Preimplantation human embryos and embryonic stem cells show comparable expression of stage-specific embryonic antigens. Stem Cells 2002; 20:329–37

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Figure 4.1 Human blastocyst displaying grade 1 expansion; no thinning of the zona pellucida and greater than 50% of the total volume filled with fluid. Clear morphological differences can be observed between the inner cell mass (center) and the trophectoderm (circling the blastocoele)

Figure 4.2 Fully expanded human blastocyst (grade 3), with thin zona

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pellucida and fully filled blastocoele. The rounded/oval inner cell mass (ICM; 6-o’clock) is made up of a group of highly compacted cells. The trophectoderm (TM) is cohesive and comprised of many cells. Cells of the inner cell mass are rounded while trophectoderm cells are elliptical and polarized

Figure 4.3 The anterior-posterior (A– P) polar axis and the dorsal-ventral axis in 2-cell and 4-cell human preembryos. (a)The second polar body marks the anterior pole and the orientation of the polar axis which determines the first cleavage plane of the zygote, while the posterior pole is opposite (A–P axis); (b) the second polar body marks the polar axis and first cleavage plane, while the dorsalventral axis marks the second cleavage plane. The second cleavage is

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equatorial and perpendicular to the first cleavage

Figure 4.4 The anterior-posterior (A– P) polar axis in the mouse pronuclear oocyte and mouse 2-cell stage embryo. (a) Pronuclear polarity is indicated by pronuclear alignment along the polar axis. F=female pronucleus in close proximity to the polar body; M =male pronucleus; (b) the second polar body marks the polar axis and first cleavage plane of the two-cell mouse embryo

Figure 4.5 The anterior-posterior (A– P) polar axis in the human pronuclear

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oocyte and human 2-cell stage preembryo. (a) Polarized prezygote with pronuclear alignment along the polar axis. Pronuclei can rotate and become organized in a polarized formation; (b) the second polar body indicates the first cleavage plane and marks the polar axis throughout development to the blastocyst stage

Figure 4.6 The anteriorposterior axes (A–P) and dorsal-ventral axes (D–V) in the mouse morula and mouse blastocyst. (a) The second polar body (PB) marks the A–P axis (reflects the A–P axis of the zygote). The D–V axis is perpendicular to the A–P axis; (b) the blastocoele occupies the abembryonic (AB) pole while the inner cell mass occupies the embryonic (EB) pole

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Figure 4.7 The anteriorposterior axes (A–P) and dorsal-ventral axes (D–V) in the human morula and blastocyst. (a) The establishment of axes in the human morula and blastocyst is less clear since one observes central ICM formation and a surrounding blastocoele; (b) in the expanded human blastocyst, the ICM is associated with one pole of the blastocyst, the embryonic pole

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Figure 4.8 Models of cell allocation and differentiation in mammalian blastocyst. The ‘Inside-outside’ hypothesis is based on the localization of individual blastomeres within cleaved preembryos and the existence of different environments surrounding inner and outer blastomeres. Blastomeres inside the preembryo form the ICM while those outside form trophectoderm. The ‘Polarization’ hypothesis is based on the polarization of each individual

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blastomere, where subsequent divisions generate daughter cells that exhibit different cytoplasmic compositions and different developmental fates. The ‘Cleavage-driven’ hypothesis is based on the early asymmetric cleavage of the preembryo. The blastomere that cleaves earlier, at the 2-cell stage, is predisposed to contribute to embryonic (ICM) cells, while slower-cleaving blastomere gives rise to abembryonic (TM) cells

Figure 4.9 Human preembryo at the compacted morula stage on day 4. This preembryo entered compaction at the 6–8-cell stage and further cell divisions were inhibited until compaction was complete. Blastomere membranes are closely apposed; several nuclei and the two polar bodies are identified easily

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Figure 4.10 Compacted human morula on day 4. Similar to the example in Figure 4.9, this morula did not complete further cell divisions until after compaction was finished and cavitation had begun. Nine to ten nuclei with a varying number of nucleoli are clearly visible

Figure 4.11 Cavitating morula on late day 4. Cells inside are ICM cells, which are round in shape and are

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loosely aggregated. Cells located peripherally are trophectoderm cells, which show polarized alignment, with one pole facing the zona pellucida while the other faces blastocoele. Cell number begins increasing once again just before the onset of cavitation

Figure 4.12 Blastocyst-stage conceptus with dominant inner cell mass and trophectoderm cells. The ICM has become smaller and extremely compacted, and forms a rounded shape. Trophectoderm cells become smaller in size with each division and take on a more elliptical shape due to increased pressure from the enlarging blastocoele

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Figure 4.13 Morphology of an inner cell mass isolated from a human day-7 blastocyst. The ICM was subsequently cultured in standard stage II sequential medium. ICM isolation involves removal of the zona pellucida with pronase followed by sequential incubation in both mouse anti-human serum and complement. (a) The isolated ICM shows an intact, compacted structure; (b) the same ICM after 2 hours of culture, showing partial decompaction and initiation of membrane blebbing; (c) 6 hours postisolation showing membrane blebbing and vesiculation with some cell lysing and possible fragmentation. This ICM likely lost its pluripotency

Figure 4.14 Visualization of cell nuclei in a living zona-free human

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blastocyst stained with Hoechst 33342. Photograph taken under double exposure using differential interference contrast (DIC) for the living image, and under UV light to demonstrate blue-stained nuclei

Figure 4.15 Oct4 expression in an expanded human blastocyst. (a) Oct4 expression in the nuclei of the ICM (green); trophectoderm is negative; (b) the same blastocyst showing ICMspecific expression of Oct4 (green) while trophectoderm nuclei stain negatively (blue only), double exposure. Cell nuclei stain blue with Hoechst 33342; Oct4+ nuclei stain aqua (overlapping greenand bluestained nuclei)

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Figure 4.16 Expression of the SSEA-1 in living human oocytes, preembryos, and blastocysts. (a) Binovular oocyte: lower oocyte was at MII of maturation at collection, was fertilized by ICSI, and produced a fragmented preembryo on day 3; upper oocyte was at Pl of maturation at collection, and never matured. The lower preembryo membranes stain positively for SSEA1 (green) by fluorescein isothiocyanate (FITC), while the upper immature

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oocyte stains negatively; (b) double exposure (DIC/FITC) of the same binovular oocyte. The lower preembryo expresses SSEA-1 (red); (c) SSEA-1 is expressed in mature oocytes (green) while immature oocytes fail to stain; (d) spatial membrane expression of SSEA-1 in a human preembryo (green); (e) SSEA-1 expression in a human morula after zona pellucida removal; (f) uniform expression of SSEA-1 in a human blastocyst; both the ICM and TM stain positively

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Figure 4.17 Expression of SSEA-3 in living human oocytes, preembryos, and blastocysts. (a)–(c) SSEA-3 in an immature human oocyte; (a) DIC; (b) SSEA-3 antibody marked with phycoerythrin R (PE) (red); (c) DIC/PE; (d)–(f) SSEA-3 in blastomeres of a human cleaved preembryo; (d) DIC; (e) SSEA-3-PE (red); (f) SSEA-3-PE (red)/DAPI

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(nuclei-blue); (g)–(k) SSEA-3 in a human blastocyst; (g) DIC; (h) DIC/DAPI (nuclei-blue); (i) SSEA-3PE in ICM cell membranes only (other cells, possibly undifferentiated TM cells or separated ICM cells showed positive staining); (j) DIC/SSEA-3-PE; (k) SSEA-3-PE/DAPI

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Figure 4.18 Expression of SSEA-4 in living human oocytes, preembryos, and blastocysts. (a) and (b) SSEA-4 in human oocytes and preembryos; (a) DIC; (b) SSEA-4 antibody marked with FITC (green). Mature oocytes and

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preembryos exhibited positive staining for SSEA-4 while the degenerating immature oocyte showed only faint positive staining; (c) SSEA-4 expression in a mature human oocyte with two polar bodies; (d) and (e) SSEA-4 in a human morula; (d) SSEA4-FITC expressed in the membranes of the inner cells only (future ICM cells?); (e) SSEA-4-FITC/DIC showing expression in inner cells, which supports the ‘lnside-outside’ hypothesis of cell allocation for blastocyst development; (f)–(i) SSEA4 expression in a human blastocyst; (f) SSEA-4-FITC staining of ICM only; (g) SSEA-4-FITC (green)/DAPI (nuclei stained blue); (h) triple staining: SSEA-4-FITC (red due to overlapping colors)/DAPI (blue nuclei)/DIC; (i) SSEA-4-FITC staining (green) of ICM only in human blastocyst

5 Human blastocysts in vitro Approximately 24 h after the morula forms, intercellular spaces begin to enlarge to create a central fluid-filled cavity called the blastocoel. The cells of the developing human blastocyst form a spherical shell enclosing the blastocoel, with one pole distinguished by a thicker accumulation of cells that makes up the inner cell mass (Figure 5.1). Although the pluripotent cells of the inner cell mass can form every type of cell found in the human body, they cannot form an entire organism since they are unable to give rise to the placenta and supporting tissues necessary for development in the human uterus. The outer ring of cells of the blastocyst constitutes the trophectoderm (Figure 5.2). While the inner cell mass gives rise to the actual embryo, the trophectoderm gives rise to the placenta and other supporting structures of pregnancy. In vivo fertilized blastocysts have been examined after being washed from the uterine cavity. Using uterine lavage performed 5 days after the luteinizing hormone peak, John Buster and his colleagues collected 25 specimens from five fertile donors who had undergone artificial insemination1,2. After examination, all recovered preembryos were transferred to recipients and resulted in three intrauterine and one tubal pregnancy. Morphological development of the specimens ranged from degenerating oocytes to mature blastocysts and all intrauterine pregnancies were established with blastocysts. A 12-cell conceptus with degenerating blastomeres was associated with the tubal pregnancy. It is of extreme interest to note that not all oocytes and preembryos collected from these donors were viable, nor were they all morphologically attractive. This perhaps points to the innate inefficiency of the human reproductive process which results in only one of four exposures to intercourse ending in a viable pregnancy. Blastocysts are generally observed in vitro sometime after 100 h post-insemination. If left in culture for 5 or 6 days after insemination, 26–65% of preembryos will reach this stage, depending on culture methods and medium composition3–7. Most in vitro fertilization (IVF) programs choose to transfer blastocysts after 5 days of culture, but the occasional clinic opts for transfer on later days, either waiting for further development of slowly growing blastocysts or delaying transfer until hatching occurs8,9. Early and late stages in blastocyst development can be recognized by both the thickness of the zona pellucida and the overall size of the blastocyst. In early blastocyst formation, the zona is thick and the diameter of the conceptus has changed little; one observes the blastocoel filling with fluid (Figure 5.3). At late stages, the blastocyst increases in overall size from the accumulated fluid, and a single ring of trophoblastic (trophectoderm) cells surrounds the cavity; the inner cell mass is clearly delineated and the zona has stretched very thin, to a mere outline (Figure 5.4). It may exist in this expanded state for 24 h or more before hatching from the zona pellucida10 (see Chapter 7). In culture, the human blastocyst can be seen to expand, collapse, and re-expand during

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late stages. Once the blastocyst has shed its glycoprotein layer, it is free to implant in the uterine wall. Of interest is that the blastocyst appears to have a built-in time clock. It has been observed that a cavity will often begin coalescing in the day-4 or -5 preembryo regardless of the number of cells present, or even after one-quarter of its cells have been removed at an earlier stage for biopsy purposes. It has been debated whether this time clock is related to chronological age, associated with the number of nuclear divisions that have taken place, or both. Cleaving preembryos have been treated experimentally with chemical agents that interrupt the cell cycle by disrupting microtubules and microfilaments. Although cell division stops under these conditions, nuclear division continues. Once exposure to the drug is discontinued, compaction and blastocyst formation occur, although fewer cells than normal are present and many of these are multinucleated11. In related experiments, 4-cell mouse preembryos were treated with cytochalasin-D to inhibit cytokinesis. When cytochalasin-D was removed, preembryos were capable of forming ‘blastocysts’ at the same time as controls, but contained only four polyploid cells. Although they possessed the ability to hatch and implant after intrauterine transfer, none were instrumental in producing living young12. It is now generally accepted, and may well be true, that the speed of cleavage is less important than the number of cells that eventually make up the blastocyst. In 1954, Hertig and colleagues recorded counts of 158 and 107 cells in two blastocysts collected at the time of hysterectomy13. In 1972, Croxatto and associates estimated 186 cells in a blastocyst obtained after uterine flushing14. Today, either we can speculate that multinucleated cells accounted for some excessive distortion of cell number in these earlier reports, or we are forced to recognize that the culturing systems used for early in vitro investigations did not produce the same cell numbers that actually occur in vivo. In 1989, nuclear fluorescent staining was used to obtain an average cell count in nine in vitro-derived blastocysts. In this study, only 58±8.1 cells were counted, with a wide range of 24–9015. Similarly, based on the results of examining 57 human specimens, Winston and colleagues estimated that two-thirds of cavitated preembryos on day 5 possessed fewer than 32 cells, and that these were multinucleated at high rates16. These last authors speculated that the majority of preembryos developing in vitro fail to complete sufficient cell cycles to produce blastocysts with cell numbers adequate to allow for normal differentiation of an inner cell mass. In the mouse, there is indeed some evidence to suggest that more rapidly dividing blastomeres contribute preferentially to the inner cell mass of the blastocyst17 (see Chapter 4). Clearly, history has demonstrated the need to improve culturing systems and further define accurate measurements to determine viability in the conceptuses we so carefully nurture in our laboratories. Coculture techniques were once believed to be necessary for encouraging preembryos to reach blastocyst stages of development. Whether coculture cells themselves impart factors favorable to support growth, or whether they utilize unnecessary medium components and/or absorb toxins, is not clearly defined. Perhaps the cells are helpful for each of these reasons, especially under conditions where culture media or environmental factors are suboptimal. In the 1990s, an occasional study showed no differences in results obtained after coculture versus conventional culture18, but most publications demonstrated enhanced rates of development after placing preembryos on various celltype monolayers, most notably Vero® cells (African green monkey kidney cells),

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autologous endometrial cells, autologous granulosa and cumulus cells, or bovine oviductal cells19–26. However, in the past few years, some of the original proponents of coculture techniques have openly declared the better suitability of sequential medium systems for extended culture27. As we will see emerging throughout this discussion, a true need exists for a marker of preembryo viability, one that will serve to guide embryologists in determining precisely what should be replaced to enhance pregnancy without exposing the patient to high-order multiple gestation. It has been proposed that one approach might involve the transfer of a single, or at most two, healthy blastocysts28. Reviewing the literature, one finds that there are reports of blastocyst implantation rates that do not surpass29, marginally surpass23 and greatly surpass30 the implantation rates of day-2 and -3 transferred conceptuses (for more detail, see Chapter 6). In the article by Gardner and colleagues30, blastocysts implanted in utero at rates in excess of 46% in a defined patient population (mean age 34.5 years, mean basal follicle stimulating hormone (FSH) 6.9 IU/l, normal uterus, no contraindication to pregnancy), representing a two-fold increase over rates in matched controls (eight patients in study group; 15 patients in control group). The culturing condition, specifically the sequential culture medium used in this study, appears to play a large role in encouraging the continued growth of viable conceptuses to advanced stages (see Chapter 2). Indeed, high rates of blastocyst development have been attained after exposing preembryos to tailored culture medium components according to the precise stage of development31. The prospect of using non-invasive techniques to select appropriate preembryos and blastocysts for transfer is a relatively new and intriguing area of investigation. It has been suggested that, in the near future, human blastocysts may be studied using ultramicrofluorescence techniques to select those with the greatest potential for implantation. In the mouse, excellent correlation was found between blastocysts with a high glucose uptake/low lactate production (as an estimate of glycolytic activity) and fetal development32. Even more recently, the concept of non-invasive amino acid profiling has caught the attention of the scientific community, since the procedure has the exciting potential to select developmentally competent single human preembryos for transfer33. The application of these and other new methods may revolutionize laboratory techniques in the future.

Influence of maternal age on successful outcome after blastocyst transfer In one study of 300 women between the ages of 18 and 45 years undergoing blastocyst transfer, the impact of maternal age was studied34. Fertilized oocytes were cultured for up to 144 h, and subsequently transferred when at least one attained an expanded blastocyst stage. The rate of cycle cancellation before oocyte retrieval increased significantly with age, and the average number of oocytes per retrieval and the proportion of cycles with expanded blastocysts declined significantly. While pregnancy rates per stimulation declined with age, pregnancy rates per transfer were approximately 50% across the entire age range. In this study, the decline in female fertility with age was concluded to be the result of reduced numbers of oocytes and the inability of fertilized oocytes to develop to

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the blastocyst stage. Implantation and pregnancy rates appeared to be unaffected by advancing age when blastocysts actually formed, a finding also observed by other investigators35. Other authors have surmised that biological ovarian age, rather than chronological age, determines blastocyst development potential and subsequent pregnancy rates28. At Cornell, we have observed a similar trend towards higher clinical pregnancy rates in selected patients suitable for day-5 transfer over the age of 40. Nonetheless, the miscarriage rate, as reflected by a lower ongoing pregnancy rate in these women, is still higher than in younger women undergoing blastocyst transfer (Tables 5.1 and 5.2). Similarly, Pantos and colleagues36 found that blastocyst development rates, clinical pregnancy rates and implantation rates were reduced across the board in women over age 40 receiving blastocysts for transfer on day 5 or 6 (22.2%, 21.1% and 8.9%, respectively vs. 40.5%, 44.6% and 19.9% in younger controls). In addition, clinical miscarriage rates were significantly higher in the older age group (25% vs. 13% in younger controls).

Monozygotic twinning and blastocyst transfer Monozygotic (MZ) twinning results from the division of a single fertilized oocyte into two genetically identical preembryos, and is thought to occur in 0.42% of all deliveries37. Several studies have shown an increased risk for MZ twinning following day-5 transfer. Rates per clinical pregnancy range from 0%38 to 12.5%39. Most reports are intermediate to these numbers, Behr and associates reporting an incidence of 5.0% per day-5 pregnancy40 and da Costa and associates reporting an incidence of 3.9%41. The Cornell program has experienced an MZ twinning rate per clinical pregnancy of 0.49% for day-3 transfer (not significantly different from natural conception) and 3.4% per day-5 transfer (significantly higher than natural conception). While MZ twins are thought to result from the division of a single fertilized oocyte to form two genetically identical preembryos, the precise mechanism(s) responsible for this division are not known. However, several observations have yielded theoretical explanations of the process. For example, the occurrence of inner cell mass ‘splitting’ might cause duplication of the preembryo at an early developmental stage (Figures 5.5 and 5.6)42,43. This splitting hypothesis was supported by animal studies showing that mechanical cleavage of mammalian preembryos in vitro could produce identical twins44,45. While these microsurgical approaches proved effective in the laboratory, the de novo fission apparently required for spontaneous MZ twinning has never been directly observed. It was very recently proposed that increased levels of glucose in culture media may predispose the inner cell mass to apoptotic changes, and if apoptotic cells are polarized in a linear fashion, cells of the inner cell mass may separate during hatching46.

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Table 5.1 Results from all day-5 transfers, with and without actual blastocysts transferred Age Number of Blastocysts (years) patients transferred/ transfer (n)

Clinical Ongoing Multiple pregnancy/ pregnancy/ pregnancy/ transfer (%) transfer (%) transfer (%)

Sacs/ blastocyst transferred (%)

<30

67

1.99±0.12

72

69

37

59

30–33

104

2.04±0.27

81

74

46

65

34–36

75

2.04±0.26

68

57

32

53

37–39

53

2.11±0.46

59

49

28

45

≥40

29

2.62±0.67

66

41

17

37

Total

328

2.09±0.38

71

62

36

55

Table 5.2 Results from all transfer cycles involving at least one true blastocyst Age(years) Number of Blastocysts Clinical Ongoing Multiple Sacs/ patients ‘transferred/ pregnancy/ pregnancy/ pregnancy/ blastocyst transfer (n) transfer (%) transfer (%) transfer (%) transferred (%) <30 30–33 34–36 37–39 ≥40 Total

64 91 71 47 26 299

1.98±0.12 2.00±0.21 2.03±0.24 2.09±0.40 2.65±0.62 2.07±0.35

75 86 70 62 69 75

72 78 61 51 46 66

38 52 34 30 19 38

61 71 56 48 39 58

The timing of twinning can be inferred from the structure of the placenta, membranes and yolk sac(s). The earlier is the twinning event, the greater is the degree to which each embryo is provided with adequate extraembryonic structures. Twin embryos that are formed early are therefore more autonomous and physiologically independent than those resulting from later twinning events47. Talansky and Gordon proposed that zona drilling experiments facilitated twinning by inducing a conformational change in some mouse blastocysts48. Specifically, some blastocoels associated with micromanipulation assumed a ‘figure of eight’ shape as the cells attempted to squeeze through the artificially created hole in the zona pellucida. Other investigators found murine blastocoel expansion unimpeded after natural hatching in most cases42, but observed trapping 5 days post-manipulation in others that received zona drilling (59/132, or 45%). One human blastocyst experimentally treated with partial zona dissection in this study hatched partially on day 8 and was noted to ‘fold double and split’, resulting in two distinctly separate but grossly unequal blastocoels. However, this connection between assisted hatching and MZ twinning remains highly speculative, as

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blastocoels and inner cell masses have never been observed to divide evenly and completely after passing through an artificial zona opening, either in humans or in any animal model. A review of six cases of MZ twins either with naturally thin zonae or where zona micromanipulation was performed suggested that zona architecture plays an important role in the development of MZ twins49. Other investigators reported five cases of MZ twins after assisted hatching from 142 pregnancies, and concluded that monoamniotic multiple gestations were increased in zona-manipulated cycles50. Recently, Schachter and co-workers reported an overall increase in MZ twinning with assisted reproductive treatment, but concluded that this increase is irrespective of treatment modality or micromanipulative techniques51. The relevance of MZ twinning in view of assisted conception outcomes is based on the markedly increased hazards attendant to pregnancies of this type. Twin-twin transfusion syndrome52,53 and fetal entanglement/umbilical cord accidents54, and other developmental anomalies55, are much more common among MZ twin gestations than in singleton pregnancies56. The subjects of MZ twinning and infertility treatments converged soon after the realization that IVF could be associated with an increased frequency of MZ twinning57. This observation was refined by a study of more than 2500 multiple births58, which postulated that ovulation induction itself, not culture conditions, was responsible for the markedly higher rate of MZ twinning following infertility treatments. Increased numbers of monoamniotic multiple gestations following zona-manipulated cycles have been reported50, but this was based on self-reported, anonymous data supplied by 42 IVF centers in the USA. More recently, a 12-fold increase in MZ twins (chorionicity not specified) was identified in a series of more than 600 patients who received single preembryo transfer59. An analysis of factors affecting zona characteristics is appropriate, since the zona pellucida is central to several theories regarding MZ twinning. A synthesis of findings from earlier studies49,57,58 suggests that at least three factors are influential in MZ twinning among patients receiving infertility treatments: ovulation induction, certain IVF culture conditions and zona architecture/micromanipulation. With these three variables occurring together so often in modern clinical infertility practice, multiple regression analysis to determine the specifie contribution of each intervention has proved difficult to perform. If the natural rate of MZ twinning (0.42%) may be considered valid in settings of spontaneous conception and single implantation, then the increase in the observed MZ twinning rate after blastocyst transfer may be partially explained by the increased number of implantations. Importantly the clinical rarity of MZ twinning challenges the study of this phenomenon in the context of IVF, as relevant investigations require large samples (>10000 cases) to detect meaningful differences with suitable statistical power60. Figures 5.7–5.10 depict day-5 blastocysts, which, after transfer to the uterine cavity generated MZ twins. Outcomes are noted.

Male/female sex ratios and birth weight after blastocyst transfer It has been postulated by several researchers that Y-bearing preembryos (males) develop at faster rates than X-bearing ones (females). If this theory is proved accurate, one might

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expect a skew in sex ratios after day-5 transfer, as the most advanced blastocysts are almost always replaced preferentially. In one very interesting study, the number of cells and metabolic activity of male and female human preembryos were examined to determine whether male preembryos were more advanced than female ones following in vitro fertilization61. The metabolic activity of normally fertilized preembryos was assessed daily by non-invasive measurements of pyruvate and glucose uptake and lactate production between days 2 and 6 after insemination. On day 6, the numbers of nuclei from the trophectoderm and inner cell masses of blastocysts were counted by differential labeling and fluorescence microscopy. Nuclei were then recovered, and the sex of the preembryos identified using nested primers to amplify the amelogenin gene and pseudogene sequences on the X and Y chromosomes, respectively. Development of male and female preembryos were then compared. From 69 of 178 (39%) preembryos that developed to the blastocyst stage, the sex of 57 was determined: 21 (37%) were male and 36 (63%) female. The number of cells in male preembryos was significantly greater on day 2 (p<0.005), and this difference was maintained through the blastocyst stage (in both the trophectoderm and the inner cell mass). Pyruvate uptake was significantly higher by male preembryos between days 2 and 5 (p<0.05). Glucose uptake and lactate production were significantly higher in male preembryos on days 4–5 (p<0.05). Extrapolation from differences in the number of cells indicated that female preembryos were approximately 4.5 h delayed in their development from day 2 onwards, compared to male preembryos. Although many thousands of children must be born before the statistical relevance of this finding can be proved, it is interesting to examine smaller numbers. In 1999, Menezo and colleagues62 reported that blastocysts transferred on day 5 after development in either coculture or sequential media systems gave rise to the birth of more male offspring, namely 225 males versus 158 females (59% vs. 41%). Contrary to this finding, replacement of thawed blastocysts did not demonstrate more males, perhaps consistent with the fact that those which were frozen actually developed more slowly to the blastocyst stage as compared to ones transferred fresh. In a report from Australia, no statistically significant shift in the male/female birth ratio was observed after blastocyst replacement, but the numbers were small and a trend towards more males was similarly observed (92 males versus 71 females or 56% vs. 44%)8. In this last study, transfers involved blastocysts which were also transferred on days 6 and 7, possibly diluting the impact of rapid cleavage. The male/female live birth ratio for the USA and the world is 1.05, or 51% males versus 49% females (UN Population Division; http:/esa.un.org/unpp [panel 2]). In our program, day-3 transfer has resulted in an expected male/female ratio of 51:49. The male/female ratio for day-5 transfer in our program is 56:44, an intriguing ‘mini-skew’ that will be followed closely as numbers of children born after blastocyst transfer continue to grow. No differences have been observed in the birth weights of live-born infants conceived through blastocyst transfer8,62, a finding dissimilar to that reported for domestic animals63.

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Grading systems used for blastocysts Different grading systems have been proposed for scoring human blastocysts. One of the most popular was developed by David Gardner and his colleagues in the late 1990s64. With this system, blastocysts are given a numerical score from 1 to 6 on the basis of their degree of expansion and hatching status as follows: 1, an early blastocyst with a blastocoel that is less than half of the volume of the embryo; 2, a blastocyst with a blastocoel that is half of or greater than half of the volume of the embryo; 3, a full blastocyst with a blastocoel completely filling the embryo; 4, an expanded blastocyst with a blastocoel volume larger than that of the early embryo, with a thinning zona; 5, a hatching blastocyst with the trophectoderm starting to herniate though the zona; and 6, a hatched blastocyst, in which the blastocyst has completely escaped from the zona. For blastocysts graded as 3–6 (i.e. full blastocysts onwards), the development of the inner cell mass is assessed as follows: A, tightly packed, many cells; B, loosely grouped, several cells; or C, very few cells. The trophectoderm is also assessed as follows: A, many cells forming a cohesive epithelium; B, few cells forming a loose epithelium; or C, very few large cells. Using this scheme, it was shown clearly that clinical pregnancy rates in excess of 60% could be attained by the transfer of at least one high-scoring blastocyst greater than or equal to 3AA38. In early 2000, we modified Gardner’s three-part grading system to fit the needs of the Cornell program. A blastocyst is defined as having a blastocoel filling greater than half the volume of the conceptus, and early stages must possess cells that suggest the formation of an inner cell mass; cavitating morulae possess smaller blastocoels, and developing inner cell masses are unidentifiable. Our current system is detailed in Figures 5.11a and 5.11b. Briefly, blastocysts are similarly given a numerical score from 1 to 6 on the basis of their degree of expansion and hatching status, but with slight modifications to Gardner’s methods: 1, an early blastocyst (blastocoel filling greater than half of the volume of the conceptus), but without overall increase in size as compared to earlier stages; 2, a true blastocyst (blastocoel filling greater than half of the volume of the conceptus) with slight expansion in overall size and some thinning of the zona pellucida; 3, a full blastocyst (blastocoel filling greater than half of the volume of the conceptus) with overall size fully enlarged and a very thin zona pellucida; 4, a hatching blastocyst (no biopsy, assisted hatching, or other major zona-manipulation); 5, a fully hatched blastocyst, completely removed from the zona pellucida (no biopsy, assisted hatching or other major zona-manipulation); and 6, a hatching or hatched blastocyst resulting from manipulations that have created a substantial hole in the zona pellucida. For blastocysts graded as 1–6 (i.e. any defined blastocyst), the development of the inner cell mass is assessed as follows: A, tightly packed, compacted cells; B, larger, loosely grouped cells or formation of a cellular bridge; C, no inner cell mass distinguishable; or D, cells of the inner cell mass appear degenerative. The trophectoderm is assessed as follows: A, many healthy cells forming a cohesive epithelium; B, few, but healthy cells, large in size, forming a loose epithelium; C, unhealthy, very large or unevenly distributed cells, may appear as few cells squeezed to the side; or D, cells appear degenerative. Using this system, transfers with at least one 1BD blastocyst (any real blastocyst by definition; 91% of cycles undergoing extended culture) result in a 75% clinical pregnancy rate and a 59% implantation rate. Transfers with at least one 3AA, 3AB or 3BA

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blastocyst result in a 78% clinical pregnancy rate and 63% implantation rate. The fact that there is so little difference between cycles with ‘early’ and late’ blastocysts indicate that any blastocyst on day 5 will lead to good pregnancy and implantation results. Only cycles without defined blastocysts on day 5 demonstrate significantly lower potential, although pregnancy and implantation are by no means negated (31% clinical pregnancy and 17% implantation). Individual grading parameters Little attention has been given to individual grading parameters (blastocyst expansion, inner cell mass quality or trophectoderm quality) as separate and individual parameters. At Cornell, we analyzed blastocysts from 156 patients on day 565. Non-hatching blastocysts were assessed for blastocoel expansion, inner cell mass morphology and trophectoderm morphology, as described above. Only blastocysts with known implantation results were included in the study. In general, there was positive correlation between overall blastocyst quality and implantation rate (for example, transfer of a 3AA blastocyst approached a 70% implantation rate). Examining individual parameters, there were no significant differences between degree of blastocyst expansion and implantation rate (3=67%, 2 =58% and 1=53%, a slight trend) or inner cell mass compaction and implantation rate (A=68%, B=62% and C=61%, a slight trend). In contrast, significantly higher implantation rates were achieved by transferring blastocysts with grade A trophectoderm (76%) compared to B (56%; p<0.05) or C (50%; p<0.05). While the morphology of the blastocyst measured with a three-part grading system impacted upon overall implantation, the quality (number and size of cells) of the trophectoderm influenced implantation to the greatest degree as a single parameter; expansion and inner cell mass morphology appeared to impact upon implantation to a lesser degree. The trophectoderm is probably the most important individual factor for implantation, as it plays a crucial role in blastocyst attachment, trophoblast development and subsequent uterine invasion. Following implantation, a good quality inner cell mass will direct embryo development. Optimal inner cell mass size and shape In contrast, a recent publication discredits the previously described grading systems as being vague and poorly defined66. The authors state that the observed differences using such systems simply reflect differences in developmental timing events rather than differences in actual quality. They further contend that combining multiple assessment factors into a single grading system without first demonstrating the importance of each factor independently is counterproductive and potentially misleading. They go on to propose alternative markers of blastocyst quality, namely the quantitative measurement of inner cell mass (ICM) size and shape, quantitative measurement of blastocyst expansion and quantitative analysis of trophectoderm cell number, first assessing the predictive value of each measure independently, then combining features with significant predictive value to form a unified grading system. What they discovered was quite interesting: the ICMs of implanting blastocysts were significantly larger than of those failing to implant (5023 µm2 vs. 4312 µn2, p=0.008). Logistic regression analysis

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revealed an approximately linear positive relationship between ICM size and implantation (p=0.01). Implantation rates ranged from approximately 20% for 2000-µm2 ICMs to nearly 50% for ICMs of 6000 µm2. A continuous model such as this is likely to reflect an accurate description of the relationship between ICM size and implantation potential. However, for simplification, they subsequently divided blastocysts into ‘large’ and ‘small’ ICM categories using a break-point that maximized the difference between the two groups. Implantation was significantly higher among blastocysts with ICM measurements >4500µm2 compared with those having smaller ICMs (45% vs. 23%, p=0.006). Optimal ICM size was therefore defined as ‘large’, measuring >4500 µm2. Inner cell masses measuring <3800 µm2 were associated with particularly low implantation rates (18%, p= 0.0028 vs. large ICMs). Blastocysts with ICMs falling between these large and small size ranges implanted at an intermediate rate of 32%, which was statistically indistinguishable from the rates for either smaller or larger ICMs. The authors failed to detect any statistically significant difference in ICM shape between implanting and non-implanting preembryos, but noted that implantation rates were highest for blastocysts with slightly oval-shaped ICMs. Blastocysts were then grouped according to whether their ICMs fell within the optimal size and/or shape ranges. ‘Topquality’ blastocysts with ICMs within both optimal size and optimal shape ranges implanted at a rate of 60% (Figure 5.12). Implantation rates were much lower for those with ICMs that were optimally sized only (29%), optimally shaped only (32%) or suboptimally sized and shaped (19%). The implantation rate for unexpanded blastocysts (10%) was significantly lower than that of high-scoring blastocysts, but no differences were seen in the mean diameter of implanting and non-implanting blastocysts that were expanded at the time of transfer (195 µm vs. 194 µm, p=0.81). The number of trophectoderm cells observed in a crosssectional circumference plane was also nearly identical between implanting and nonimplanting conceptuses (11.0 vs. 10.8, p=0.64). From this study, it was concluded that quantitative measurements of the inner cell mass are highly indicative of implantation potential, and that these measurements provide more predictive value for implantation when compared to the qualitative assessment of developmental timing events. Whether or not this conclusion is accurate will be determined only after other investigators apply these methods, replicate the published results and determine the basis for the correlative findings.

Blastocyst development and intracytoplasmic sperm injection It was suggested in 1994 that blastocyst development rates may be reduced when oocytes are fertilized by sperm derived from suboptimal semen samples67. Since that time, other investigators have reported lower blastocyst formation rates associated with intracytoplasmic sperm injection (ICSI) procedures using suboptimal sperm when compared to non-ICSI treatment68–71. Possiblehypotheses for impaired blastocyst formation after ICSI have ranged from defective sperm DNA and delayed male pronucleus formation to abnormal male centrosome control through the first cleavages and inappropriate or delayed genomic activation.

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Extended culture has even been proposed as a means of weeding out chromosomal abnormalities contributed by defective spermatozoa. This idea has resulted from the concern that the use of ICSI may contribute to increased numbers of abnormal offspring72,73. The relationship between poor preembryo quality and the existence of low morphological ‘normalcy’ of a semen sample has certainly been suggested74. It has been proposed that, by using a selection process such as extended culture to identify preembryos most likely to develop normally, chances of an errant paternal genome being inherited by ICSI might be reduced75. While an interesting speculation, recent studies suggest that the phenotypic manifestation of paternal genomic abnormalities probably does not occur before implantation and, therefore, blastocyst transfer may not diminish the likelihood of inheriting genetic defects involving ‘male factor’ loci76. Experience in the Cornell program indicates that blastocyst development is not impaired after ICSI procedures are carried out. Although slightly poorer development after ICSI initially occurred in our first year of day-5 transfer trials, rates of clinical pregnancy and implantation have since evened out for conventional insemination and ICSI groups, both being in the region of 71% and 55%, respectively, for all preembryos transferred, actual blastocysts or not.

Enzymatic removal of the zona pellucida before blastocyst transfer It is theorized that the zona pellucida is required during early preembryonic development, at the 2-cell to 8-cell stages, but may not be necessary aiter compaction occurs. The first report of an ongoing human pregnancy following the intrauterine transfer of day-6 zonafree blastocysts was presented in early 199777. In this case report, a patient had undergone eight previous transfers without success; preembryos were cocultured on Vero® cells until blastocyst stages, and their zonae removed with 0.5% pronase. Clinical pregnancy was established after the transfer of two zona-free blastocysts, and led to an ongoing singleton gestation. In a follow-up report by the same authors published in 1998, 19 additional women (mean age 32.6 years, and 2.1 previous failed attempts) had zona-free blastocysts transferred78. Clinical pregnancy and implantation rates were 53% and 33%, respectively. The multiple pregnancy rate was 40%. Based on these studies, it appears that zona-manipulated blastocysts implant relatively well. The authors suggest that enzymatic treatment of the zona may allow better anchorage and dialog of the blastocyst with the endometrium. Other investigators have reported similar outcomes after the transfer of zona-free blastocysts79.

Blastocyst development after manipulation of the zona pellucida Artificially created holes, cutsor flaps are sometimes made purposefully in the zonae pellucidae of selected preembryos. This occurs routinely during biopsy assisted hatching and ICSI procedures. These breeches, if large enough, may affect subsequent blastocyst expansion and hatching events (see Chapter 7). Blastocyst development after biopsy for preimplantation genetic diagnosis differs considerably from what is routinely observed in non-manipulated conceptuses. Because a relatively large hole is created in the zona

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pellucida during the biopsy procedure, either with chemical agents or ‘burned’ with a laser, premature hatching often occurs without coincident expansion of the blastocyst or gradual thinning of the zona pellucida. Cavitating morulae and early non-expanded blastocysts are often observed to herniate through the artificially made holes in the zona pellucida (Figures 5.13–5.15). Preembryos subjected to assisted hatching procedures follow much the same course after blastocysts form if chemical agents are used to breech the zona (Figures 5.16 and 5.17), whereas assisted hatching techniques utilizing a mechanical tearing action are associated with slits or flaps rather than actual holes, and have been implicated in blastocyst entrapment during expansion and hatching. Small punctures made with an ICSI pipette during assisted fertilization procedures are likely to either reseal or be too tiny to permit easy escape of the expanding blastocyst.

Uterine receptivity and blastocyst transfer In the rabbit, preembryos have been shown to implant at greater rates as the interval between ovulation and transfer increases. This finding has been attributed to the transfer being carried out concurrent to higher circulating progesterone levels in the mother. It has been put forth that this state tends to inhibit uterine contractions80. It was subsequently determined that uterine junctional zone contractions progressively decrease as the cycle moves into the luteal phase, and that fewer contractions are associated with improved pregnancy rates81. It has additionally been demonstrated that day 5 is favorable for the human in terms of decreased uterine contractility82.

Time line for optimal blastocyst development Based on studies performed at Cornell, examining a timely first cleavage and the importance of observing growth to the 8-cell stage by the morning of day 383, we worked out a model for optimal blastocyst development (Table 5.3). Other factors could certainly be incorporated into the time line such as standards for pronculear size and alignment, morphological features felt to be of importance during cleavage stages or inner cell/trophectoderm characteristics, but these would require that preembryos be ‘perfect’ rather than simply ‘timely’. Briefly, the time line covers the two major points listed above and sets up minimal growth parameters for assessing adequate blastocyst development by the time of intrauterine transfer (Figure 5.18). Figures 5.19–5.23 depict groups of preembryos that were serially photographed on days 3, 4 and 5 of development (preembryo/morula/blastocyst stage). Some closely follow the proposed time line, others do not. Outcomes are noted. Figure 5.24 depicts sequential photographs of mouse preembryos developing to the hatching blastocyst stage.

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Blastocysts known to implant Figures 5.25–5.35 present blastocysts on day 5 of culture that are known to have implanted after transfer to the uterine cavity. Each led to a fetal heartbeat by ultrasound, and most often produced a living, healthy child. Where this is not the case, the obstetrical outcome is indicated in the legend.

Favorable blastocysts that failed to implant Figure 5.36 shows highly graded blastocysts that failed to implant after intrauterine transfer.

Unfavorable development on day 5 with implantation Figure 5.37 demonstrates poorly graded day 5 conceptuses that led to ongoing pregnancies after transfer.

Table 5.3 Time line for optimal blastocyst development Time line

Actual time

Hour 0

15.00, day 0 insemination or ICSI

Before hour 24 (ICSI)

15.00, day 1 cleavage to the 2-cell stage, ≤10% fragments

Before hour 26 (insemination)

17.00, day 1 cleavage to the 2-cell stage, ≤10% fragments

Before hour 42

09.00, day 2 cleavage to the 4-cell stage, ≤10% fragments

Before hour 66

09.00, day 3 cleavage to the 8-cell stage, ≤10% fragments

Before hour 72

15.00, day 3 evidence of increased cell-cell contact

Before hour 90

09.00, day 4 evidence of uniform compaction

Before hour 100

19.00, day 4 evidence of central cavitation

Before hour 114

09.00, day 5 grade 2 blastocyst expansion

Before hour 120

15.00, day 5 transfer; grade 3 blastocyst expansion

Before hour 144

15.00, day 6 if not transferred, evidence of hatching or already hatched

ICSI, intracytoplasmic sperm injection

Minimal expected observation

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Abnormal blastocyst development Blastocysts can form despite possessing discernable inner cell masses (Figure 5.38), with very small inner cell masses (Figure 5.39) or with abnormally developed inner cell masses (Figure 5.40). They may also form despite having exhibited extensive vacuolization during development fig. 5.41). Cells constituting the trophectoderm may also display abnormal features (Figure 5.42). Recall that, although the pluripotent cells of the inner cell mass can form every type of cell found in the human body, they cannot form an entire organism since they are unable to give rise to the placenta and supporting tissues necessary for development in the human uterus. While the inner cell mass gives rise to the actual embryo, the trophectoderm gives rise to the placenta and other supporting structures of pregnancy. It has been observed that some blastocysts possess non-viable trophectoderm cells despite exhibiting healthy-appearing inner cell masses. In such specimens, the trophectoderm eventually degenerates, while the inner cell mass proliferates and enlarges (Figure 5.43). It has been proposed that because these specimens are incapable of establishing a viable pregnancy, their inner cell masses might be useful, without ethical objection, for potentially life-saving therapies resulting from appropriately designed stem cell research.

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57. Edwards RG, Mettler L, Walters DE. Identical twins and in vitro fertilization. J In Vitro Fertilization Embryo Transf 1986; 3:114–17 58. Derom C, Vlietinck R, Derom R, Van den Berghe H, Thiery M. Increased monozygotic twinning rate after ovulation induction. Lancet 1987; 1:1236–8 59. Blickstein I, Verhoeven HC, Keith LG. Zygotic splitting after assisted reproduction. N Engl J Med 1999; 340:738–9 60. Sills ES, Moomjy M, Zaninovic N, et al. Human zona pellucida micromanipulation and monozygotic twinning frequency after IVF. Hum Reprod 2000; 15:890–5 61. Ray PF, Conaghan J, Winston RM, Handyside AH. Increased number of cells and metabolic activity in male human preimplantation embryos following in vitro fertilization. J Reprod Fertil 1995; 104:165–71 62. Menezo YJ, Chouteau J, Torello J, Girard A, Veiga A. Birth weight and sex ratio after transfer at the blastocyst stage in humans. Fertil Steril 1999; 72:221–4 63. McEvoy TG, Sinclair KD, Young LE, Wilmut I, Robinson JJ. Large offspring syndrome and other consequences of ruminant embryo culture in vitro: relevance to blastocyst culture in human ART. Hum Fertil (Camb) 2000:3:238–46 64. Gardner DK, Schoolcraft WB. In vitro culture of the human blastocyst. In Jansen R, Mortimer D, eds. Towards Reproductive Certainty: Infertility and Genetics Beyond 1999. Carnforth, UK: Parthenon Publishing, 1999:378–88 65. Zaninovic N, Berrios R, Clarke RN, Bodine R, Ye Z, Veeck LL. Blastocyst expansion, inner cell mass (ICM) formation, and trophectoderm (TM) quality: is one more important for implantation? Presented at the 57th Annual Meeting of the American Society for Reproductive Medicine, Orlando, FL, October 2001 66. Richter KS, Harris DC, Daneshmand ST, Shapiro BS. Quantitative grading of a human blastocyst: optimal inner cell mass size and shape. Fertil Steril 2001; 76:1157–67 67. Janny L, Menezo YJ. Evidence for a strong paternal effect on human preimplantation embryo development and blastocyst formation. Mol Reprod Dev 1994; 38:36–42 68. Jones GM, Trounson AO, Lolatgis N, Wood C. Factors affecting the success of human blastocyst development and pregnancy following in vitro fertilization and embryo transfer. Fertil Steril 1998; 70:1022–9 69. Miller JE, Smith TT. The effect of intracytoplasmic sperm injection and semen parameters on blastocyst development in vitro. Hum Reprod 2001; 16:918–24 70. Shoukir Y, Chardonnens D, Campana A, Sakkas D. Blastocyst development from supernumerary embryos after intracytoplasmic sperm injection: a paternal influence? Hum Reprod 1998; 13:1632–7 71. Wun WS, Wun CC, Valdes CT, Dunn RC, Grunert GM. Blastocyst formation is a good indicator for attainment of assisted reproduction. Chin J Physiol 1997; 40:237–42 72. Bonduelle M, Joris H, Hofmans K, Liebaers I, Van Steirteghem A. Mental development of 201 ICSI children at 2 years of age. Lancet 1998; 351:1553 73. Bowen JR, Gibson FL, Leslie Gl, Saunders DM. Medical and developmental outcome at 1 year for children conceived by intracytoplasmic sperm injection. Lancet 1998; 351:1529–34 74. Ombelet W, Fourie FL, Vandeput H, et al. Teratozoospermia and in-vitro fertilization: a randomized prospective study. Hum Reprod 1994; 9:1479–84 75. Sakkas D. The use of blastocyst culture to avoid inheritance of an abnormal paternal genome after ICSI. Hum Reprod 1999; 14:4–5 76. Banerjee S, Lamond S, McMahon A, Campbell S, Nargund G. Does blastocyst culture eliminate paternal chromosomal defects and select good embryos?: inheritance of an abnormal paternal genome following ICSI. Hum Reprod 2000; 15:2455–9 77. Fong CY, Bongso A, Ng SC, Anandakumar C, Trounson A, Ratnam S. Ongoing norrmal pregnancy after transfer of zona-free blastocysts: implications for embryo transfer in the human. Hum Reprod 1997; 12:557–60

An atlas of human blastocysts 140 78. Fong CY, Bongso A, Ng SC, Kumar J, Trounson A, Ratnam S. Blastocyst transfer after enzymatic treatment of the zona pellucida: improving in-vitro fertilization and understanding implantation. Hum Reprod 1998;13:2926–32 79. Jones GM, Trounson AO, Gardner DK, Kausche A, Lolatgis N, Wood C. Evolution of a culture protocol for successful blastocyst development and pregnancy. Hum Reprod 1998; 13:169–77 80. Adams CE. Retention and development of eggs transferred to the uterus at various times after ovulation in the rabbit. J Reprod Fertil 1980; 60:309–15 81. Fanchin R, Righini C, Olivennes F, Taylor S, de Ziegler D, Frydman R. Uterine contractions at the time of embryo transfer alter pregnancy rates after in-vitro fertilization. Hum Reprod 1998; 13:1968–74 82. Fanchin R, Ayoubi JM, Righini C, Olivennes F, Schonauer LM, Frydman R. Uterine contractility decreases at the time of blastocyst transfers. Hum Reprod 2001; 16:1115–19 83. Zaninovic N, Veeck L, Clarke RN, Rosenwaks Z. Early assessment of human preembryos as an indicator for potential blastocyst development. Presented at the 56th Annual Meeting of the American Society for Reproductive Medicine, San Diego, CA, October 2000

Figure 5.1 Schematic: early and late blastocyst stages

Figure 5.2 Trophectoderm (TM). (a) Large individual TM cells; inner cell mass (ICM) forms a band that spans the blastocoele; (b) smaller individual TM cells as compared to (a) and compacted ICM; insert focuses on TM cells; (c) on high magnification, one observes quite large TM cells that appear pushed to one side; a cytoplasmic ‘string’ can be seen between separating cells. According to Scott (Semin Reprod Med 2000; 18:171), the persistence of these processes after full expansion of the blastocyst correlates negatively with subsequent implantation; (d) high

magnification of a cytoplasmic ‘string’ between developing ICM and TM

Figure 5.3 Blastocoelic enlargement. To the left, an early cavitating morula; to the right, an expanded blastocyst. Note the enlarging blastocoele and thinning zona pellucida as development advances

Figure 5.4 The zona pellucida thins to a mere outline as the blastocoele enlarges

Figure 5.5 Inner cell mass trapping as hatching occurs through a small hole. Note part of the ICM inside the zona pellucida and part outside, which could, theoretically, lead to ICM ‘splitting’

Figure 5.6 Developing blastocyst that appears to possess two separately cavitating segments

Figure 5.7 Day-5 transfers that led to monozygotic twins. (a) Two transferred blastocysts that led to two gestational sacs and three fetal hearts by ultrasound investigation. The patient elected to selectively reduce the monozygotic twins and one healthy female child was ultimately delivered; patient aged 36 years; (b) two blastocysts replaced; two sacs/three fetal hearts by ultrasound; natural reduction; one healthy male child delivered; patient aged 38 years

Figure 5.8 Day-5 transfers that led to monozygotic twins. (a) Two blastocysts replaced; two identical males and one female delivered; patient aged 27 years; (b) two blastocysts replaced; two identical males and third male sibling delivered; patient aged 36 years

Figure 5.9 Day-5 transfers that led to monozygotic twins. (a) Two blastocysts replaced; two gestational sacs/three fetal hearts noted by ultrasound investigation. The monozygotic twins were miscarried; one female delivered to this oocyte recipient; patient aged 45 years; (b) two blastocysts replaced; two gestational sacs/three fetal hearts noted by ultrasound investigation. The monozygotic twins were reduced; one female delivered; patient aged 31 years

Figure 5.10 Day-5 transfer that led to monozygotic twins. Two blastocysts replaced; two gestational sacs/three fetal hearts noted by ultrasound investigation. The monozygotic twins were reduced; one male delivered; patient aged 42 years

Figure 5.11 Blastocyst grading schemes used by the Cornell program. (a) Grading system detailed in text and

photograph; (b) grading system detailed by individual photographs (see next page)

Figure 5.12 Blastocysts graded for ‘top-quality’ inner cell masses. (a) Large, compacted, and oval inner cell masses; both blastocysts implanted and two females were delivered; (b) large, compacted, and oval inner cell masses; one of these blastocysts implanted and a male child was delivered

Figure 5.13 Cavitating morula/early blastocyst herniating through artificially-made hole in zona pellucida. A hole was made on day 3 during biopsy for preimplantation genetic diagnosis; the conceptus was ultimately diagnosed as being monosomic for chromosomes 15 and 22

Figure 5.14 Early blastocyst, diagnosed for trisomy 13 after PGD, seen here herniating through the artificially made hole in the zona pellucida

Figure 5.15 Early blastocyst, diagnosed for trisomy 13 and trisomy 21 after PGD, seen here herniating through artificially made hole in zona pellucida

Figure 5.16 Non-expanded blastocyst after PGD, herniating through biopsy site in zona pellucida

Figure 5.17 Late blastocyst after PGD, herniating through artificially made hole in zona pellucida

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Figure 5.18 Time line for optimal blastocyst development

Figure 5.19 Preembryos serially photographed on days 3, 4, and 5 of development (preembryo/morula/blastocyst stages); patient aged 32 years. (a)–(c) Two preembryos photographed over 3 days and transferred on day 5. Neither

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implanted; (d)–(f) five preembryos from the same patient, photographed over 3 days and cryopreserved on day 5. Because the patient did not become pregnant after fresh transfer, she returned to have these blastocysts thawed. Three were thawed and replaced; an ongoing pregnancy with one fetal heart is currently in progress beyond 20 weeks

Figure 5.20 Preembryos serially photographed on days 3, 4, and 5 of development (preembryo/morula/blastocyst stages);

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patient aged 32 years. (a)–(c) Three preembryos photographed over 3 days, two of which were transferred on day 5 (the one on the left and the one on the right in photograph (c). An ongoing pregnancy with one fetal heart is currently in progress beyond 20 weeks

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Figure 5.21 Preembryos serially photographed on days 3, 4, and 5 of development (preembryo/morula/blastocyst stages);

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patient aged 29 years. (a)–(c) Three preembryos photographed over 3 days, one of which was transferred on day 5 (middle blastocyst in photograph (c) along with a second blastocyst not shown. A pregnancy with two fetal hearts was established which is ongoing beyond 20 weeks

Figure 5.22 Preembryos serially photographed on days 3, 4, and 5 of development

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(preembryo/morula/blastocyst stages); patient aged 29 years. (a)–(c) Four preembryos photographed over 3 days, two of which were transferred on day 5 (the two blastocysts to the left in photograph (c). Both implanted and a twin pregnancy is ongoing beyond 20 weeks

Figure 5.23 Individual preembryos serially photographed on days 3, 4, and 5. (a) On the afternoon of day 3, cellcell aggregation (earliest evidence of impending compaction) is noted; on day 4, cavitation begins to occur

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between still large blastomeres; on day 5, the blastocyst possesses a round and compacted inner cell mass despite having a trophectoderm that is made up of few, large cells. This conceptus was frozen for future replacement in an oocyte recipient patient; no thaw has yet been carried out on this 36-year-old woman; (b) on the afternoon of day 3, ten blastomeres are noted; on day 4, an early blastocyst with poor inner cell mass definition is apparent; on day 5, the conceptus has developed into a good late blastocyst with a cohesive and multicellular trophectoderm, a slightly small inner cell mass, and possesses an extremely thin zona pellucida. This blastocyst was replaced along with one other that was less well developed (grade 2BB) and a pregnancy that started out with two gestational sacs was established; an ongoing singleton pregnancy with one fetal heart is now beyond 20 weeks; patient aged 40 years; (c) this conceptus, photographed on days 3, 4, and 5, possesses an abnormal zona pellucida. On day 3, a healthy appearing 12-cell preembryo was observed; on day 4, it had developed into a healthy appearing cavitating morula; by day 5, a blastocyst of mediocre quality has formed despite the persisting zona defect. This blastocyst was replaced, but pregnancy was not established. The patient shown here has undergone nine separate transfers between the ages of 24 and 31 years of age, twice with blastocysts,

only achieving pregnancy one time with day-3 preembryos

Figure 5.2 (a)–(i) Sequential photographs of mouse morulae developing to the hatching blastocyst stage

Figure 5.25 Day-5 blastocysts known to implant after their replacement. (a) One healthy female child delivered; maternal age 31 years; (b) two healthy female children delivered; maternal age 35 years; (c) two healthy male children delivered; maternal age 30 years; (d) two healthy female children delivered; maternal age 35 years

Figure 5.26 Day-5 blastocysts known to implant after their replacement. (a) Two healthy male children delivered; maternal age 28 years; (b) two healthy male children delivered; maternal age 38 years; (c) two healthy male children delivered; maternal age 33 years; (d) two healthy male children delivered; maternal age 30 years

Figure 5.27 Day-5 blastocysts known to implant after their replacement. (a)

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One healthy male and one healthy female delivered; maternal age 31 years; (b) one healthy male and one healthy female delivered; maternal age 28 years; (c) one male and one female delivered prematurely; both babies died soon after birth; maternal age 32 years; (d) one healthy male and one healthy female delivered; maternal age 35 years

Figure 5.28 Day-5 blastocysts known to implant after their replacement. (a) Two healthy male children delivered; maternal age 30 years; (b) two healthy male children delivered; maternal age 35 years; (c) one healthy male and one healthy female delivered; maternal age 31 years; (d) two healthy male children delivered; maternal age 34 years

Figure 5.29 Day-5 blastocysts known to implant after their replacement. (a) Two healthy male children delivered; maternal age 38 years; (b) one healthy male and one healthy female delivered; maternal age 30 years; (c) two healthy female children delivered; maternal age 34 years; (d) two healthy female children delivered; maternal age 35 years

Figure 5.30 Day-5 blastocysts known to implant after their replacement. (a) Two healthy male children delivered;

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maternal age 30 years; (b) two healthy male children delivered; maternal age 31 years; (c) two healthy male children delivered; maternal age 32 years; (d) two healthy male children delivered; maternal age 38 years

Figure 5.31 Day-5 blastocysts known to implant after their replacement. (a) One healthy male and one healthy female delivered; maternal age 32 years; (b) one healthy male and one healthy female delivered; maternal age 33 years; (c) two healthy males delivered; maternal age 35 years; (d) one healthy male and one healthy female delivered; maternal age 35 years

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Figure 5.32 Day-5 blastocysts known to implant after their replacement. (a) One healthy male and one healthy female delivered; maternal age 29 years; (b) two fetal heart pregnancy; one healthy male ultimately delivered; maternal age 25 years; (c) two healthy females delivered; maternal age 42 years (oocyte donation); (d) two healthy females delivered; maternal age 45 years (oocyte donation)

Figure 5.33 Day-5 blastocysts known to implant after their replacement. (a) Two healthy males delivered; maternal

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age 38 years; (b) two healthy females delivered; maternal age 29 years; (c) two fetal heart pregnancy ongoing greater than 30 weeks; maternal age 53 years (oocyte donation); (d) two fetal heart pregnancy ongoing greater than 30 weeks; maternal age 32 years (oocyte donation)

Figure 5.34 Day-5 blastocysts known to implant after their replacement. (a) Two fetal heart pregnancy ongoing greater than 20 weeks; maternal age 29 years; (b) two fetal heart pregnancy ongoing greater than 20 weeks; maternal age 33 years; (c) two fetal heart pregnancy ongoing greater than 20 weeks; maternal age 31 years; (d) two fetal heart pregnancy ongoing greater than 12 weeks; maternal age 31 years

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Figure 5.35 Day-5 blastocysts known to implant after PGD. (a) Ongoing singleton pregnancy greater than 20 weeks; maternal age 38 years; (b) ongoing singleton pregnancy greater than 12 weeks; maternal age 33 years; (c) ongoing singleton pregnancy greater than 12 weeks; maternal age 41 years; (d) ongoing twin pregnancy greater than 12 weeks; maternal age 38 years

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Figure 5.36 Highly graded blastocysts that failed to implant. (a) Two blastocysts that failed to implant after transfer to the uterine cavity; patient age 33 years; (b) two blastocysts that failed to implant after transfer to the uterine cavity; patient age 36 years; (c) two blastocysts that failed to implant after transfer to the uterine cavity; patient age 29 years; (d) blastocyst exhibiting a zona pellucida that is conjoined to the zona of a degenerative oocyte. After separation of the two, the healthy appearing blastocyst was transferred but failed to implant; patient age 36 years

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Figure 5.37 Poorly graded day-5 conceptuses that led to ongoing pregnancy. (a) These compacting morulae were transferred on day 5 to a 33-year-old woman; one implanted and a healthy male child was subsequently delivered; (b) three cavitating morulae on day 5 which were replaced in a 42year-old woman and led to the birth of a healthy female child; (c) three morulae that were transferred on day 5 to a 47-year-old oocyte donation recipient and gave rise to two gestational sacs by ultrasound; ultimately, one healthy female child was delivered

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Figure 5.38 (a)–(d) Blastocysts without visible inner cell masses

Figure 5.39 (a) and (b) Blastocysts with small inner cell masses

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Figure 5.40 Blastocysts with apparently abnormal inner cell masses. (a) A blastocyst that appears to possess two inner cell masses at 1 o’clock and 9 o’clock positions; (b) a blastocyst that appears to possess two inner cell masses at 12 o’clock and 4 o’clock, plus a third thickening of TM cells at 7 o’clock; (c) a blastocyst exhibiting a completely degenerative inner cell mass; (d) high magnification of a blastocyst with three large cells surrounding a small, compacted inner cell mass; the two inserts to the right show the same blastocyst using different focal planes on lower magnification

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Figure 5.41 Sequential photographs of vacuolated conceptuses on days 3,4, and 5 of development

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(preembryo/morula/blastocyst stages). (a)–(c) No vacuolization was noted in any of the three preembryos on day 3 of development. On day 4, the upper left morula exhibited extensive vacuolization. On day 5, the same conceptus (upper left) developed into a relatively healthy appearing expanded blastocyst; note small vacuoles in cells of the TM. The patient did not become pregnant after replacement of the uppermiddle blastocyst. Same patient as in Figure 5.23 (c) who failed to achieve pregnancy in eight of nine transfer attempts

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Figure 5.42 Abnormal trophectoderm. (a) Appears to have one large cell pushed to the upper and right sides; (b) single, unhealthy appearing cell pushed to the upper and right sides; (c) few cells make up this thin and unhealthy appearing trophectoderm; (d) while generally healthy in appearance, the trophectoderm is

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comprised of few cells; (e) despite a developing inner cell mass, the trophectoderm of this conceptus is made up of few cells; (f) this highly interesting blastocyst appears to have a second ring of TM cells within its blastocoele

Figure 5.43 (a)–(d) Degenerating trophectoderm associated with inner cell mass proliferation and enlargement

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6 Preembryo selection and blastocyst quality: how to choose the optimal conceptus for transfer Reducing the incidence of multiple pregnancy Through the application of extended culture techniques, we have been given the opportunity to offer more responsible medical care to our patients by reducing the number of preembryos for transfer. Unfortunately, selective reduction of one or more fetuses is all too often necessary when higher-order gestations are established, and the procedure itself carries obvious medical and emotional risks1–4. Multifetal gestations are the largest single cause of poor obstetrical outcome and subsequent neonatal difficulties, these pregnancies being associated with increased incidences of pre-eclampsia, gestational diabetes, pregnancy-induced hypertension, preterm labor, low birth weight and extensive neonatal care (Figure 6.1)5. Patients and health-care professionals often present differing perspectives on the concept of establishing a multiple pregnancy6. When patients, embryologists and clinicians were given a questionnaire soliciting views on the potential risks and benefits of blastocyst culture and multiple pregnancy, it was discovered that patients were far more accepting of multiple pregnancy as a prospective outcome than those involved in their treatment, especially multiple pregnancy involving twins7. This greater acceptance was demonstrated despite an awareness of the concrete risks associated with multiple gestation. It is uncertain, however, how many patients would take the same view retrospective to actually experiencing a high-order gestation. It must be assumed that few would embrace the potential ordeal without reservation.

History of day-3 and day-5 transfer Until the late 1990s, preembryos were routinely cultured until only day 2 or 3 before being transferred to the uterus. The primary impedance to culturing for longer periods involved the lack of an appropriate culture medium to sustain the viability of compaction and blastocyst development. In an effort to optimize clinical pregnancy rates under those conditions, clinics felt compelled to replace up to four day-3 preembryos, or even more, in women over the age of 35 or 40 years. The downside to this replacement strategy was that too many women were placed at risk of having triplet or quadruplet gestations (Figure 6.2). Researchers then found that better than acceptable clinical pregnancy rates could be achieved with extended culture using specialized media and replacement of only two

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conceptuses that had grown to the blastocyst stage (Figure 6.3). The identification of key regulators and recognition of changing physiological requirements over the course of 5 days of preembryo growth led these researchers to refine the basic culture media used during critical days, and to develop what are now commonly referred to as ‘sequential’ media8,9. Because extended culture selects for preembryos with high implantation potential, clinical pregnancy and implantation rates after the replacement of only one or two day-5 conceptuses has been demonstrated to be quite high in selected patient populations, without the associated risk of high-order multiple births.

Advantages of day-5 transfer Additional days of culture clearly single out preembryos with high implantation potential; those likely to result in viable offspring continue developing until day 5 of culture, forming healthy-appearing blastocysts with compacted inner cell masses and proliferating trophectoderm, highly suitable for replacement (Figure 6.4). Conversely, those arresting in culture may be selectively excluded from replacement. In the Cornell program, 328 day-5 transfers carried out on selected in vitro fertilization (IVF) patients (age range 24–44 years) from late 1999 to mid-2002 yielded the following results: clinical pregnancy 71%, implantation 55%. The average number of day-5 preembryos replaced was 2.09. Recipients of donor oocytes fared even better (average age 42 years): 82% clinical pregnancy and 66% rate of implantation. Although most patients described here were selected for blastocyst transfer based on number and quality of their conceptuses on days 1–3, some were included because of the medical necessity to transfer only a single preembryo or because transfer had to be delayed, and others because they fervently desired extended culture. Those patients who selected themselves for day-5 transfer despite the laboratory’s recommendations were often those who failed in the effort. Unfortunately the transfer of two blastocysts in our program resulted in a twinning rate per pregnancy of 52%. This means that over one-half of the women undergoing day-5 transfer in an effort to reduce their chances for multiple pregnancy actually ended up with precisely what they wished to avoid. The finding of excessive rates of twinning despite a conservative approach to the number of blastocysts being transferred has been demonstrated by other investigators outside of the Cornell program10. Very similar to our results, David Gardner and colleagues described a prospective randomized trial of selected patients (good responders to ovarian stimulation) in which day-5 transfers achieved 71% pregnancy and 51% implantation with an average of 2.2 blastocysts replaced11. While the pregnancy rate for matched day-3 transfers was not significantly different (66%), the implantation rate was lower (30%), despite a higher number of preembryos being replaced per transfer (3.7). Schoolcraft and co-workers, in a multicenter trial, reported a 66% ongoing pregnancy rate and 48% implantation rate in 174 selected women undergoing day-5 transfer12. An average of 2.2 blastocysts was replaced per transfer. In a separate publication, Schoolcraft and Gardner described 229 oocyte recipients who established an 88% clinical pregnancy and a 66% implantation rate with 2.1 blastocysts replaced per transfer13. Oocyte recipients have been noted by other investigators to be particularly good

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candidates for extended culture, their rates of pregnancy and implantation exceeding those of non-donor day-5 transfers, even after controlling for donor age14. In 2000, Milki and associates reported a 68% pregnancy rate and 47% implantation rate (average number transferred 2.4), which surpassed the 46% pregnancy and 20% implantation observed in a similar patient population with transfer on day 3 (average number transferred 4.6)15. In this study, all patients were under the age of 40 and possessed at least three 8-cell preembryos on day 3. For those IVF programs attempting blastocyst transfer in every patient walking through the door, there may be evidence of clinical advantage as well. An increase was reported by Marek and colleagues in ongoing pregnancy from 36% for day-3 transfer (in about 1997) to 44% for day-5 transfer (in about 1998) in non-selected women16. Not surprisingly, implantation rates were also significantly higher for day-5 transfers (23% vs. 34%) as a result of fewer conceptuses being transferred (3.0, day 3 and 2.5, day 5). Although this retrospective analysis of clinical data involved distinctly separate time periods and the use of different culture media between groups, it clearly demonstrates no detrimental effect after applying day-5 protocols for all patients. Similarly, Wilson and associates reported an increase in clinical and ongoing pregnancy rates in unselected patients with blastocyst transfer, particularly those under the age of 35 years, but also for their program as a whole17. Unfortunately, the management of an extended culture program is not as simple as purchasing new media and delaying transfer for 2 days. Minute deficiencies in gas regulation, temperature control or other environmental conditions affecting pH or resulting in premature amino acid degradation, barely recognizable with short-term culture, may be exacerbated by prolonged culture. In addition, variability in lots of some purchased media has been shown to contribute to mutable outcomes. As a result, many programs have experienced difficulty in maintaining high pregnancy rates with blastocyst transfer, and have ultimately dropped the procedure altogether. Their frustrations are often expressed on internet mail forums and during scientific exchange at meetings.

Comparisons of day-3 and day-5 transfer Prospective and retrospective comparisons of day-3 versus day-5 results are rife with potential grouping mismatches. All too often, patients opt for one procedure or the other based upon what they have read, their previous experiences, what has happened to their friends or their desire for singleton births. As well, embryologists often convert a potential blastocyst transfer to day 3 because of concern over preembryo quality or quantity. A more appropriate means of comparing results would involve prospective application of blastocyst transfer to all even-numbered cycles and day-3 transfer to all odd-numbered cycles, and not to sway from this position until a very large number of patients had been treated without bias (same time period, same media, same incubators and environment, same embryologists). Regrettably, neither our internal review boards nor our well-educated patients would accept this sort of mandate, and we are therefore forced to draw conclusions from non-randomized populations. Comparing smaller, but similar, groups does not always take into account the differences in changing laboratory technique and personnel, preembryo fragmentation, zona pellucida characteristics,

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number of previous attempts or impact of gynecological history, uterine anomalies, transfer difficulties or other important clinical features that ultimately affect outcomes. When one seeks to review the scientific literature on the issue of day-3 versus day-5 transfer, reports are varied. Some investigators have found no differences in either clinical pregnancy or implantation rate after day-5 replacement18–24, some have noted a better implantation rate only11,25–29 and yet others have experienced a general overall improvement in implantation and pregnancy rates in patients who have had blastocyst transfer13–17,30,31. In some programs, it has been suggested that blastocyst transfer may be detrimental for women demonstrating poor preembryo quality on day 3, or at least detrimental in unselected patient populations27. Another publication suggests that pregnancy and implantation rates decline dramatically in repeated cycles of blastocyst transfer following one or more unsuccessful blastocyst attempts32. One report describes the experience of extended culture resulting in an almost 50% decrease in implantation and pregnancy rates33. At Cornell, we elected to pursue blastocyst transfer because of the disturbingly high triplet rate in good-responding patients with at least one morphologically optimal preembryo (grade 1 or 1.5) on day 3. Examination of our data identified 139 women, aged 24–39, with three preembryos replaced on day 3 who would have been automatic candidates for day-5 transfer if the technology had been available. In these cycles, the clinical pregnancy rate was 88.5%, the implantation rate was 61% and the multiple rate was 62.6%, 19% of which were triplets. Undoubtedly, transferring only two preembryos in these cycles would have proved reasonable, probably producing similar clinical pregnancy results without triplets. Now, after more than 2 years of working with blastocysts, we are seeing pregnancy and implantation rates not significantly different after day-5 transfer in the same type of patient (71% and 55%, respectively), with only two blastocysts transferred in most cases. With the confidence of these results, we instigated a protocol to transfer electively only two preembryos on day 3 for consenting patients who met criteria for blastocyst transfer. Again, this study was not completely randomized since it required patient consent rather than assignment. Nonetheless, results of elective transfer of two preembryos on day 3 looked extremely similar to two blastocysts on day 5 when patient populations were closely matched. In our first investigation, 24 patients elected to accept two, rather than three, preembryos; 17/24 established clinical pregnancy (71%), and a 56% implantation rate was realized. It is rather surprising how close these numbers are to our current day-5 transfer results. Moreover, doing a simple computer search of all patients treated during the past year under the age of 40 who elected to have two preembryos replaced on day 3, despite the fact that they had more than two available, yielded similar results. Fifty-nine patients achieved a 68% clinical pregnancy rate and a 49% implantation rate. In an effort to elucidate further the differences between day-3 and day-5 transfers in selected patient populations, we will continue to accumulate numbers for this investigation.

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Experience of the United Kingdom Other programs have published large studies finding no difference in clinical pregnancy rates when two, rather than three, preembryos were transferred34–41. In a massive analysis of outcome data from the United Kingdom, elective transfer of two preembryos was demonstrated to maintain that country’s current standards in pregnancy rates, compared favorably to having three replaced and served to reduce the multiple pregnancy rate42. The group studied risk factors in 44236 transfer cycles involving 25240 women. They examined age, cause and duration of infertility, number of previous attempts, number of previous live births, number of oocytes fertilized and number of preembryos replaced. It was determined that only previous live birth could be shown to be correlated with a significantly higher birth rate after IVF. Conversely, older age, tubal infertility, longer duration of infertility and a greater number of previous attempts was associated with significantly lower birth and multiple birth rates. Interestingly, it was shown that when more than four oocytes were fertilized, there was no difference at all in the birth rate whether two or three preembryos were replaced. However, there was a considerable increase in the multiple birth rate with three (Figure 6.5).

Developments in selection criteria for day-3 transfer The quest to look for markers of preembryo viability at early stages of development continues. If such markers can be easily identified without invasion or trauma to the conceptus, we will not need to depend on growth characteristics alone to select the lowest number of conceptuses for transfer. Pronuclear distribution and alignment Despite the fact that human preembryos are usually selected for transfer using morphology criteria on days 2 and 3, earlier morphological indicators may assist in predicting implantation and pregnancy potential. One group has examined the relationship between pronuclear morphology and subsequent blastocyst development43. In this study, prezygotes were scored according to distribution and size of nucleoli within each pronucleus (Figure 6.6). Those displaying equalities between the nuclei demonstrated 49.5% blastocyst formation, while those with unequal sizes, numbers or distribution of nucleoli formed blastocysts only 28% of the time44. Cleaving preembryos that were selected initially by zygote morphology and secondarily by morphology on day 3 demonstrated increased pregnancy and implantation (31% and 57%, respectively), compared with those selected by morphology alone (19% and 33%, respectively; p<0.01). Furthermore, the authors demonstrated that there was a significant difference between prezygote-scored and non-scored cycles on day 3 (pregnancy, 57% vs. 33%; implantation, 31% vs. 19%) and on day 5 (pregnancy, 73% vs. 58%; implantation, 52% vs. 39%). Other investigators have reported similar improvements in pregnancy after incorporating pronuclear assessment into their selection criteria45–51. In contrast, the Cornell program was unable to demonstrate the value of assessing either nucleolar distribution or alignment after extensive data were collected from more

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than 2000 fertilized oocytes52. We sought to determine whether the morphology of pronuclear-stage oocytes could be used to assess subsequent development. A retrospective evaluation of 2187 prezygotes from 258 patients was performed. Prezygotes were graded according to their pattern of nucleolar distribution: PZ1 (nucleoli in both pronuclei were aligned at pronuclear junctions); PZ2 (one pronucleus had aligned nucleoli but the second pronucleus exhibited scattered nucleoli); PZ3 (both pronuclei possessed scattered nucleoli); or PZ4 (one or both pronuclei had only 1–2 nuclei) (Figure 6.7). In addition, each prezygote was graded according to the orientation of pronuclei relative to the polar bodies: PB+ pronuclei were perpendicular to polar bodies and PBpronuclei were parallel to polar bodies (Figure 6.8). Prezygotes were later assessed for cleavage at 24 h, day-3 preembryo quality (cell number and fragmentation) and day-5 development to the blastocyst stage. Of the 2187 conceptuses assessed after fertilization, 20% were classified as PZ1, 34% PZ2, 38% PZ3 and 8% PZ4. In addition, 70% were categorized as PB+ and 30% were PB-. Few differences were observed in any subgroup according to their cleavage status at 24 h (either still pronuclear, in syngamy or at the 2cell stage). A slightly higher proportion of PZ3 zygotes cleaved by 24 h (p<0.03), while differences were not found among other subgroups. In addition, the proportion of prezygotes developing into good-quality preembryos on day 3 (8-cell stage, <20% fragmentation) was slightly higher (p=0.047) in the PZ1 subgroup as compared to the other subgroups, although the clinical relevance of this statistical difference was questionable (24% PZ1; 20% PZ2; 17% PZ3; 20% PZ4; 20% PB+; 20% PB−). Nor were any significant differences found in the capability of the various prezygote subgroups to develop to the blastocyst stage by day 5 of culture (PZ1 30%; PZ2 26%; PZ3 30%; PZ4 30%; PB+ 31%; PB− 25%). From these data, we concluded that no clinically relevant differences between any of these prezygote subgroups could be deemed beneficial for prediction of subsequent preembryo or blastocyst quality. Timeliness of first cleavage and subsequent cleavage rate Another indicator of preembryo viability involves the assessment of sperm-penetrated oocytes at 24 h after intracytoplasmic sperm injection (ICSI), or 26 h after insemination. Break-points of 24 and 26 h were chosen based on studies demonstrating that oocytes fertilized through ICSI procedures tended to undergo the first cleavage 2–3 h earlier than those produced through insemination53. It has been observed by us and others that oocytes undergoing a timely first cleavage (by 24–26 h) often produce more 8-cell conceptuses on day 3 and healthier-appearing blastocysts on day554–56. In one study from Cornell, it was demonstrated that a higher proportion of blastocysts on day 5 originated from 2-cell preembryos at 24/26 h when compared to those that were in syngamy or pronuclear stages (Figure 6.9), and the stage at 24/26 h in combination with the cleavage stage on the morning of day 3 proved extremely useful in predicting blastocyst development on day 555. It was notable that the predictive value of day 3 morphology was dependent on evaluating preembryos by about 66 h after insemination or ICSI (mornings). Delaying day-3 evaluations until 72 h (afternoons) diluted the predictive correlation of the assessment. Furthermore, fewer than eight, or more than eight, blastomeres on the morning of day 3 had a significantly negative association with normal blastocyst development, a finding closely confirmed by

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other investigators (Figure 6.10)57. While it was not surprising to note that low blastomere numbers on day 3 were suboptimal, it was extremely interesting to verify the growing suspicion that preembryos with 12 or 14 blastomeres were not particularly better because of their advanced growth. It has become apparent that preembryos continuing to cleave beyond the 8-cell stage without undergoing compaction are less likely to develop normally to the blastocyst stage. Failure to undergo the compaction process at the appropriate stage indicates that the internal ‘clock’ regulating sequential developmental events is faulty. In the course of these investigations, it was also observed that an occasional oocyte began fragmenting even before the first cleavage (Figure 6.11). When fragmentation occurred so early, severely impaired development followed. In contrast, compromised development was not noted when minor or moderate fragmentation (<20%) first occurred during or after the first cleavage. Based on the results described above, we can now predict that a fertilized oocyte exhibiting two cells at 24 h with less than 20% fragmentation, followed by eight cells on the morning of day 3, will have a very high chance (>50%) of forming a viable blastocyst by day 5. We routinely look at conceptuses at 24/26 h to assist in identifying potential elective transfers of two conceptuses on day 3, as well as identifying potential extended culture candidates. Criteria used in our program to aid in selecting patients for blastocyst transfer include at least four syngamous or cleaved preembryos at the 24/26-h investigation, at least three 8-cell or six 7–9-cell preembryos on the morning of day 3 and fragmentation levels less than 20% in all preembryos meeting these conditions. We have proposed a model, suitable for our program, that describes a time line for optimal development to the blastocyst stage (see Table 5.3 and Figure 5.18). Several other programs use slightly different criteria for selecting patients for blastocyst transfer. In the early work of Gardner and colleagues, normal basal follicle stimulating hormone (FSH) levels and ten follicles >12 mm on the day of human chorionic gonadotropin (hCG) administration were required for inclusion in day-5 protocols11. The group of Milki and Behr required at least three 8-cell preembryos on day 3 to go forward with extended culture15. Similarly, Racowsky and co-workers used eight or more fertilized prezygotes to select patients for blastocyst transfer, and concluded that a minimum of three 8-cell preembryos on day 3 was necessary in order to realize improved outcomes with blastocyst protocols27. Cytoplasmic fragmentation and multinucleation In addition to cleavage rates, Alikani and colleagues examined the degree of fragmentation on day 3, specific fragmentation pattern and blastomere multinucleation in relation to the success of blastocyst formation57. They discovered that more than 15% fragmentation on day 3 (Figure 6.12a) resulted in lower blastocyst formation on day 5 (16.5% vs. 33.3% with 1–15% fragmentation). Furthermore, type IV fragmentation (large, randomly distributed fragments, often necrotic and associated with uneven blastomeres; Figure 6.12b) led to a significant reduction in the percentage of blastocysts formed (14.7%) as compared to type I, II or III patterns (small and associated with a single blastomere, localized to the perivitelline space, or small and scattered: 38.6%, 32.9% and 32.4% respectively). The presence of multinucleated blastomeres (Figure

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6.13) also had a negative impact on further development, only 15.9% of preembryos displaying multinucleation in one or more cells giving rise to a healthy blastocyst (versus 31.9% without multinucleation). Collectively, these data suggest that early cleavage anomalies significantly reduce the ability of preembryos to proceed with normal development to days 5 and 6. Value of day-3 morphology as a single assessment Several investigators have reported a very inexact correlation between a day-3 evaluation alone and subsequent normal blastocyst development58,59 (Figures 6.14 and 6.15. A recent publication describes a study wherein the best two preembryos chosen on day 3 were cultured separately from the others for an additional 2 days60. These were preembryos that would have been transferred had replacement been carried out on day 3. On day 5, the two most advanced blastocysts were chosen for transfer. Examining associations between the choices on day 3 and the blastocysts chosen for transfer on day 5 revealed that both choices from day 3 were transferred only 23% of the time. At least one of those chosen on day 3 was transferred 38% of the time and 39% of the cycles had neither choice transferred. However, after extending culture to day 6, it was noted that 68% of the ‘picks’ on day 3 ended up being either transferred on day 5 or of a quality sufficient to be frozen on day 5 or 6. Blastocyst development and chromosomal abnormalities Human preembryos cultured in vitro present high rates of chromosomal anomalies. These anomalies may be numerical (e.g. trisomies or aneuploidy) or structural (e.g. translocations). The incidence of chromosomal abnormalities in cleaved preembryos after fluorescence in situ hybridization (FISH) analysis ranges between 15 and 85%, depending on the number and specificity of probes used. A high percentage of aberrant preembryos exhibit mosaic or chaotic abnormalities61. Chromosomal anomalies are even higher in arrested preembryos, suggesting that preembryo selection may exclude some abnormalities before implantation. For this reason it has been proposed that culturing preembryos to blastocyst stages will select chromosomally normal conceptuses. Another possibility represents selective allocation of aneuploid cells to the trophectoderm. This hypothesis was questioned recently after mosaicism was demonstrated in the inner cell mass of the human blastocyst62. The overall frequency of mosaicism has been shown to increase from 15% at the 2–4-cell stage to 49% at the 5–8-cell, 58% at the morula and up to 90% at the blastocyst stage39. On the other hand, certain forms of mosaicism, e.g. chaotic mosaicism and diploid-aneuploid mosaicism, were reduced at the blastocyst stage. The highest rates of mosaicism seen in blastocysts have been reported to be 2npolyploid mosaicism, usually diploid/tetraploid63,64. It has also been shown that aneuploid cell lines from cleavage stages may persist to the blastocyst stage, but their incidence is very low65. Most polyploid cells in the blastocyst are thought to arise during blastocyst formation and are probably precursors of trophoblast differentiation66. Sandalinas and colleagues sought to correlate which chromosomal abnormalities were compatible with prolonged development in vitro67. This group took preembryos assessed as abnormal following preimplantation genetic diagnosis (PGD) analysis, allowed them

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to grow in culture until day 5 or 6 and correlated which abnormalities were associated with subsequent blastocyst formation. They found that extensive mosaicism could be detected in some blastocysts developing in vitro and that 37% of trisomic preembryos reached the blastocyst stage of development. They further reported that only monosomies X and 21, monosomies compatible with development into the first trimester, were actually found in conceptuses reaching the blastocyst stage. The authors concluded that, although there was a strong selection bias against chromosomally abnormal preembryos with extended culture, the selective potential was limited in its ability to exclude many clinically relevant aberrations, especially the trisomies. Magli and associates reported similar findings65. Fifty-nine per cent of day-3 aneuploid preembryos did not reach the morula or blastocyst stages of development in vitro. Nonetheless, 40% of blastocysts studied by FISH techniques were shown to possess aneuploid cells, and a significant number of preembryos diagnosed as aneuploid on day 3 produced blastocysts with inner cell mass mosaicism. These data suggest that euploid cells are not necessarily preferentially allocated to inner cell masses, and that aneuploid cells are not necessarily preferentially allocated to the trophectoderm of affected blastocysts. A high concordance between day-3 aneuploidy diagnosis and inner cell mass lineage was observed with trisomies (97%), while a reduced concordance was found with monosomies (65%) and haploidies (18%). Further studies will be needed to correlate the chromosomal status of the blastocyst cultured in vitro with blastocyst morphology and implantation potential. Blastocysts have been noted to form after a single pronculeus was observed at 18 h post-insemination, suggesting that this feature alone does not indicate developmental incompetence68. The Cornell group has similarly observed this type of outcome after documentation of a monopronuclear status (Figure 6.16). It has been suggested that all inseminated oocytes displaying a single pronucleus should be rechecked within 2–6 h to confirm pronuclear number. One report where investigators did this revealed that 25% of repeat observations led to the visualization of two pronuclei rather than one69. Such a finding suggests that developmental asynchrony between male and female pronuclei may not be a rare event. A single pronucleus can form and persist after in vitro insemination or sperm injection, resulting in a gynogenic or rare androgenic haploid prezygote. Human oocytes displaying a single pronucleus have been observed to continue growth to blastocyst stages, and some of these have been shown actually to be diploid. One such study describes removing the pronucleus from single-pronucleate oocytes after conventional insemination. Of 16 pronuclei examined, six were diploid, four of which contained XY chromosomes70. This suggests that sperm and oocyte nuclei can associate to form a single diploid pronucleus, as opposed to simply being asynchronous in development. It has further been shown that preembryos developing from monopronucleate oocytes after IVF insemination are most often fertilized and diploid, while those forming from activated ICSI oocytes are almost always haploid parthenotes69,71,72. For this reason, preembryos developing from inseminated oocytes displaying one pronucleus may be replaced, although not preferentially; those developing after ICSI should not be transferred. Since it has been described that diploid pronuclei are often larger than usual owing to their double amount of DNA73, this characteristic may be favorable in monopronculeate oocytes after in vitro insemination.

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When should we transfer only a single conceptus? Research from our program suggests that when a fertilized oocyte displays two cells at the 24/26-h observation, grows to an 8-cell stage by the morning of day 3 and possesses less than 20% fragmentation, it has a very high implantation potential. Add the situation where it develops into a high-grade blastocyst, and it has a greater than 70% chance of implanting in the uterine cavity (Figure 6.17). These conceptuses should probably be replaced alone, as the twinning rate is unacceptably high with two. Additionally, it has been proposed by other groups that elective transfer of a single blastocyst would benefit patients with poor obstetrical outcomes after previous high-order gestations74.

Should we be transferring blastocysts for all patients? We must first acknowledge that extended culture will not turn a poor-quality day-3 preembryo into a healthy blastocyst. The value of this technology is to provide better selection criteria when educated choices must be made. Neither will extended culture confer any special advantage to a woman with only one or two oocytes at harvest, at least not at this time. Our data show that with good-responding patients, transferring two preembryos on day 3 serves the same purpose as using blastocysts in terms of clinical pregnancy and reduction of high-order gestation. The efforts described here with our own day-3 and day5 trials, and those reported by most others, are primarily dealing with this type of patient. For programs unable to carry out blastocyst transfer, unhappy with the variability in media or results, or unwilling to extend culture to day 5, options are available for the selection of appropriate preembryos on day 3. Such programs should consider examining fertilized oocytes at 24/26 h to assess for timely cleavage, and then evaluate preembryos on the morning of day 3, keeping in mind that the predictive value for blastocyst development may be lost if one waits until the afternoon. If the first cleavage has occurred by 24 h and at least two 8-cell day-3 preembryos of good quality (< 20% fragmentation) are available, one should consider replacing only two conceptuses in women under the age of 40 years. Keep in mind that, whichever stage is chosen for carrying out transfer, a clinic’s ultimate goal should be to help produce a single, healthy baby. Transfer of one conceptus alone will effectively eliminate all possibilities for multiple gestation, excluding identical twins. As responsible investigators and care-givers, our quest for the future must follow this direction.

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24. Lundqvist M, Rova K, Simberg N, Lundkvist O. Embryo transfer after 2 or 5 days of IVF culture: a retrospective comparison. Acta Obstet Gynecol Scand 2002; 81:126–32 25. Hsieh YY, Tsai HD, Chang FC. Routine blastocyst culture and transfer: 201 patients’ experience. J Assist Reprod Genet 2000; 17:405–8 26. Plachot M, Belaisch-Allart J, Mayenga JM, Chouraqui A, Serkine AM, Tesquier L. Blastocyst stage transfer: the real benefits compared with early embryo transfer. Hum Reprod 2000; 15(Suppl 6):24–30 27. Racowsky C, Jackson KV, Cekleniak NA, Fox JH, Hornstein MD, Ginsburg ES. The number of eight-cell embryos is a key determinant for selecting day 3 or day 5 transfer. Fertil Steril 2000; 73:558–64 28. Vidaeff AC, Racowsky C, Rayburn WF. Blastocyst transfer in human in vitro fertilization. A solution to the multiple pregnancy epidemic. J Reprod Med 2000; 45:529–39, discussion 539– 40 29. Karaki RZ, Samarraie SS, Younis NA, Lahloub TM, Ibrahim MH. Blastocyst culture and transfer: a step toward improved in vitro fertilization outcome. Fertil Steril 2002; 77:114–18 30. Cruz JR, Dubey AK, Patel J, Peak D, Hartog B, Gindoff PR. Is blastocyst transfer useful as an alternative treatment for patients with multiple in vitro fertilization failures? Fertil Steril 1999; 72:218–20 31. Van Der Auwera I, Debrock S, Spiessens C, et al. A prospective randomized study: day 2 versus day 5 embryo transfer. Hum Reprod 2002; 17:1507–12 32. Shapiro BS, Richter KS, Harris DC, Daneshmand ST. Dramatic declines in implantation and pregnancy rates in patients who undergo repeated cycles of in vitro fertilization with blastocyst transfer after one or more failed attempts. Fertil Steril 2001; 76:538–42 33. Levron J, Shulman A, Bider D, Seidman D, Levin T, Dor J. A prospective randomized study comparing day 3 with blastocyst-stage embryo transfer. Fertil Steril 2002; 77:1300–1 34. Fujii S, Fukui A, Yamaguchi E, Sakamoto T, Sato S, Saito Y. Reducing multiple pregnancies by restricting the number of embryos transferred to two at the first embryo transfer attempt. Hum Reprod 1998; 13:3550–4 35. Devreker F, Emiliani S, Revelard P, Van den Bergh M, Govaerts I, Englert Y. Comparison of two elective transfer policies of two embryos to reduce multiple pregnancies without impairing pregnancy rates. Hum Reprod 1999; 14:83–9 36. Devreker F, Emiliani S, Revelard P, Govaerts I, Vannin AS, Englert Y. [Diminishing the risk of multiple pregnancies in in vitro fertilization: from selective transfer of two embryos to that of one blastocyst?] Rev Med Brux 1999; 20:A463–7 37. Matson PL, Browne J, Deakin R, Bellinge B. The transfer of two embryos instead of three to reduce the risk of multiple pregnancy: a retrospective analysis. J Assist Reprod Genet 1999; 16:1–5 38. Dean NL, Phillips SJ, Buckett WM, Biljan MM, Tan SL. Impact of reducing the number of embryos transferred from three to two in women under the age of 35 who produced three or more high-quality embryos. Fertil Steril 2000; 74:820–3 39. Ludwig M, Schopper B, Katalinic A, Sturm R, AI-Hasani S, Diedrich K. Experience with the elective transfer of two embryos under the conditions of the German embryo protection law: results of a retrospective data analysis of 2573 transfer cycles. Hum Reprod 2000; 15:319–24 40. Licciardi F, Berkeley AS, Krey L, Grifo J, Noyes N. A two-versus three-embryo transfer: the oocyte donation model. Fertil Steril 2001; 75:510–13 41. Ng EH, Lau EY, Yeung WS, Ho PC. Transfer of two embryos instead of three will not compromise pregnancy rate but will reduce multiple pregnancy rate in an assisted reproduction unit. J Obstet Gynaecol Res 2001;27:329–35 42. Templeton A, Morris JK. Reducing the risk of multiple births by transfer of two embryos after in vitro fertilization. N Engl J Med 1998; 339:573–7 43. Scott LA, Smith S. The successful use of pronuclear embryo transfers the day following oocyte retrieval. Hum Reprod 1998; 13:1003–13

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63. Bielanska M, Tan SL, Ao A. Chromosomal mosaicism throughout human preimplantation development in vitro: incidence, type, and relevance to embryo outcome. Hum Reprod 2002; 17:413–19 64. Ruangvutilert P, Delhanty JD, Serhal P, Simopoulou M, Rodeck CH, Harper JC. FISH analysis on day 5 post-insemination of human arrested and blastocyst stage embryos. Prenat Diagn 2000; 20:552–60 65. Magli MC, Jones GM, Gras L, Gianaroli L, Korman I, Trounson AO. Chromosome mosaicism in day 3 aneuploid embryos that develop to morphologically normal blastocysts in vitro. Hum Reprod 2000; 15:1781–6 66. Benkhalifa M, Janny L, Vye P, Malet P, Boucher D, Menezo Y. Assessment of polyploidy in human morulae and blastocysts using co-culture and fluorescent in-situ hybridization. Hum Reprod 1993; 8:895–902 67. Sandalinas M, Sadowy S, Alikani M, Calderon G, Cohen J, Munne S. Developmental ability of chromosomally abnormal human embryos to develop to the blastocyst stage. Hum Reprod 2001; 16:1954–8 68. Gras L, Trounson AO. Pregnancy and birth resulting from transfer of a blastocyst observed to have one pronucleus at the time of examination for fertilization. Hum Reprod 1999; 14:1869–71 69. Staessen C, Janssenswillen C, Devroey P, Van Steirteghem AC. Cytogenetic and morphological observations of single pronucleated human oocytes after in-vitro fertilization. Hum Reprod 1993; 8:221–3 70. Levron J, Munne S, Willadsen S, Rosenwaks Z, Cohen J. Male and female genomes associated in a single pronucleus in human zygotes. Biol Reprod 1995; 52:653–7 71. Palermo GD, Munne S, Colombero LT, Cohen J, Rosenwaks Z. Genetics of abnormal human fertilization. Hum Reprod 1995; 10(Suppl 1): 120–7 72. Sultan KM, Munne S, Palermo GD, Alikani M, Cohen J. Chromosomal status of uni-pronuclear human zygotes following in-vitro fertilization and intracytoplasmic sperm injection. Hum Reprod 1995; 10:132–6 73. Austin CR. The Mammalian Egg. Oxford: Blackwell Scientific Publications, 1961 74. Damario MA, Phy JL, Tummon IS. Successful elective single blastocyst transfer in a patient with prior repetitive high-order multiple gestations. J Assist Reprod Genet 2002; 19:205–8

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Figure 6.1 Ultrasound images of triplet pregnancies. Photographs courtesy of Steven Spandorfer, Weill Medical College of Cornell University. (a) Three individual sacs; (b) two sacs, three embryos (monozygotic twins)

Figure 6.2 Ultrasound image of a quadruplet pregnancy. Inserts show individual embryos. Photograph

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courtesy of Isaac Kligman, Weill Medical College of Cornell University

Figure 6.3 Human blastocysts replaced on day 5. High-graded blastocysts, each with a prominent ICM and cohesive TM

Figure 6.4 Human blastocysts on day 5. High-graded blastocysts suitable for replacement. In these cases only one blastocyst should be replaced as the implantation rate is quite high

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Figure 6.5 Odds of birth and multiple birth related to age, number of oocytes fertilized, and number of preembryos replaced (Templeton A, Morris JK. N Engl J Med 1998; 339:573). The authors studied birth and multiple birth risk factors in 44236 cycles (25240 women) by examining age, cause and duration of infertility, number of previous attempts, number of previous live births, number of oocytes fertilized, and number of preembryos

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transferred. They found that older age, tubal infertility, longer duration of infertility, and greater number of previous attempts correlated with significantly lower birth and multiple birth rates while previous live birth correlated with significantly higher birth rate (but not multiple birth rate). (a) When more than four oocytes were fertilized, there was no difference in the birth rate whether two or three preembryos were replaced. The odds of a birth were calculated as compared with those of a 30-year-old woman with more than four oocytes fertilized and two preembryos replaced (odds for such a woman, 1.0); (b) there was, however, a considerable increase in the multiple birth rate when three were replaced. The odds of a multiple birth were calculated as compared with those of a 30-year-old woman with more than four oocytes fertilized and two preembryos replaced (odds for such a woman, 1.0)

Figure 6.6 Nucleolar distribution in human male and female pronuclei approximately 18 hours postinsemination. (a) Nucleoli are nearly equal in number and size and are

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closely aligned at pronuclear junctions, an example of a good pronuclear score; (b) unequal number and size of nucleoli with scattered distribution; a poor pronuclear score

Figure 6.7 Pronuclear grading system according to pattern of nucleolar distribution. (a) PZ1: both pronuclei have aligned nucleoli; (b) PZ2: one pronucleus with aligned nucleoli, the other exhibiting scattered nucleoli; (c) PZ3: both pronuclei possess scattered nucleoli; (d) PZ4: few nucleoli in one or both pronuclei (in this instance, both)

Figure 6.8 Orientation of pronuclei in relation to polar body (PB) position. (a) PB+: pronuclei are perpendicular to PB, in correct position for syngamy and the first cleavage. (b) PB-: pronuclei are parallel to PB, necessitating rotation before syngamy

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Figure 6.9 Assessment of preembryo development at 24/26 hours postICSI/insemination. (a) At 24 hours post-injection, this prezygote possesses two pronuclei (2PN); (b) at 26 hours post-insemination, no PN are seen in this syngamous oocyte; (c) 2-cell conceptus at 24 hours after ICSI; (d) graph depicting how blastocyst development on day 5 correlates to the number of blastomeres observed on the morning (am) of day 3. The 8-cell stage, on the morning of day 3, is optimal for blastocyst development on

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day 5. The majority of 8-cell preembryos that develop into blastocysts on day 5 arise from those that possessed two cells at 24/26 hours (approximately 56%). It is interesting to note that preembryos with more than eight cells on the morning of day 3 form blastocysts at lower rates than those with exactly eight cells

Figure 6.10 Predictive criteria for day5 blastocyst development were lost if preembryos were not evaluated until the afternoon of day 3. Some conceptuses with fewer than eight cells in the morning developed to eight cells by the afternoon (pm), yet their ability to form blastocysts was impaired

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Figure 6.11 Fragmentation before the first cleavage. (a) and (b) Pronuclear oocytes with excessive fragmentation 24 hours after injection. Such fragments are not reincorporated into the zygote and developmental arrest generally follows

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Figure 6.12 (a) Day-3 preembryo exhibiting two to three blastomeres surrounded by multiple cytoplasmic fragments. Due to the reduction of available cytoplasm for cleavage, these are usually unable to develop further; (b) type IV fragmentation (per Alikani et al.58), with large, randomly distributed fragments and associated with unevenly sized and shaped blastomeres; these are associated with a significant decrease in subsequent blastocyst formation

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Figure 6.13 Multinucleation. (a) Multinucleated blastomere in a day-2 2-cell preembryo. The blastomere to the left possesses three nuclei; there are two nuclei in the blastomere to the right (overlapping from this angle). Multinucleation of all blastomeres in a 2-cell conceptus is associated with impaired blastocyst development; (b) multinucleated blastomere in a day-2 4-cell preembryo. The blastomere to the left presents with at least two nuclei. There is less negative impact on subsequent development when multinucleation first occurs in a single blastomere in preembryos possessing four or more cells

Figure 6.14 (a) and (b) Preembryos with good morphology on day 3 that

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subsequently failed to form viable blastocysts

Figure 6.15 (a) and (b) Day-3 preembryos exhibiting good morphology, but which failed to reach the blastocyst stage of development after continued culture. Each possesses 8–10 blastomeres and none are associated with more than very minor cytoplasmic fragmentation. Unfortunately, correlation between day-3 morphology and subsequent blastocyst development is not precise

Figure 6.16 Human blastocyst on day 5, developed from a prezygote

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displaying one pronucleus (1PN) after insemination. The ICM is welldeveloped and compacting; the TM is made up of large, but healthy, cells

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Figure 6.17 A 2-cell conceptus at 24/26 hours that grows to eight cells by the morning of day 3 and then subsequently develops into a healthy appearing blastocyst, which possesses an extremely high implantation potential, exceeding 70%. Single blastocyst replacement is recommended under these growth and development conditions. (a) Two-cell stage at 24 hours after ICSI; (b) eight blastomeres on the morning of day 3; (c) high-graded blastocyst on day 5

7 Blastocyst hatching The zona pellucida is a glycoprotein layer that surrounds oocytes and preembryos during the preimplantation period. A number of authors have demonstrated that the human zona is composed of three acidic glycoproteins called ZP1, ZP2 and ZP31–3. The zona has various possible roles in preimplantation development: prevention of polyspermy, protecting the preembryo from physical or immunological damage and preserving its integrity during cleavage. Although the zona is sometimes described as having roles in sperm binding, in induction of the acrosome reaction and as a matrix for sperm penetration, these last phenomena may be the simple consequence of its presence around the oocyte (personal communication, Michael Bedford). After compaction and formation of cell junctions, this protective coat is no longer necessary for the developing human preembryo, at least under in vitro culture conditions. Hatching is a process that involves blastocyst escape through the zona pellucida, a prerequisite for normal implantation. In the human, implantation is initiated by trophoblastic invasion of the uterine endometrium. As the blastocyst expands, there is a gradual accumulation of fluid in the blastocoel, resulting in increased pressure on both trophectoderm and the zona pellucida. At the same time, cells of the trophectoderm proliferate rapidly to form a cohesive monolayer4. It is not entirely clear whether the concomitant thinning of the zona pellucida results from physical pressures exerted by blastocyst expansion and trophectoderm proliferation, or whether thinning is primarily the effect of lytic enzymes produced by the developing preembryo, or both5. There is some evidence to support the involvement of enzymes, since viable preembryos demonstrate very gradual thinning of the zona pellucida, or at least show variations in zona thickness, even before blastocyst expansion6. In contrast, arrested preembryos do not7. Because the overall size of the preembryo does not increase until cavitation, it can be deduced that at least some aspects of zona pellucida thinning are related to enzymes released from the preembryo. While the process of hatching is well characterized in the mouse, it is less well understood in the human. It is possible that some or all of the mechanisms involved are similar. In the mouse, the in vitro hatching mechanism involves, at the least, a trypsinlike proteinase, called strypsin8. Before hatching, strypsin is found in mural trophectoderm cells, presumably at the site where hatching will occur. After hatching, strypsin is again identified around the opening from which the blastocyst emerges. But proof of trypsin-like protease involvement in mouse hatching comes from studies of inhibitors. Yamazaki and colleagues showed that hatching was suppressed in 83% of murine embryos by a synthetic trypsin inhibitor with no adverse effect on subsequent blastocyst development9. A potential substrate for the trypsin-like protease action may be ZP310. Experiments utilizing an antihatching mouse model support the theory of embryoproduced enzyme involvement11. Furthermore, in vivo hatching studies also reveal zona lysis activity during the peri-implantation period. It is possible that non-embryonic

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uterine enzymes are at least partly responsible for the complete shedding of the zona pellucida in vivo12. At the onset of hatching, the mouse blastocyst generates greater than ten times the amount of superoxide anion radicals than in the pre- and post-hatching stages. In fact, adding superoxides to cultured zonae induces thinning and hatching within a matter of seconds, the result of superoxides disrupting the glycosilic bonds of zona glycoproteins13. Interestingly, hatching can occur in the absence of an inner cell mass, supporting the primary importance of the trophectoderm for this process11,14,15. In the mouse, hatching occurs in areas adjacent to mural trophectoderm opposite to the inner cell mass16. Although more variable in the human, the hatching site usually develops relatively close to the inner cell mass in proximity to the polar trophectoderm (Figures 7.1–7.4). In human blastocysts cultured in vitro, small membrane blebs or vesicles frequently protrude through the zona pellucida before hatching occurs (Figures 7.5 and 7.6). Contrary to popular belief, they do not always indicate the precise location of subsequent hatching. After the zona pellucida has begun to rupture (Figure 7.7), the ultimate size of the opening created is roughly between one-quarter and one-third of the circumference of the zona pellucida, permitting easy and rapid blastocyst escape (Figures.7.8 and 7.9). Once an opening is created, the blastocyst begins to protrude through the zona. After this, several mechanisms may be involved in the completion of hatching. If lytic enzymes and blastocoelic tension are primarily responsible for zona pellucida rupture, then another mechanism must be responsible for complete extrusion and escape of the blastocyst. It has been suggested that actin polymerization plays a role in this regard, since actin filaments are heavily concentrated on trophectoderm cells, especially so at the site of hatching. The initial protrusion is probably mediated through continuous actin polymerization of trophectoderm cell microfilaments. When cytochalasin-B, an inhibitor of actin polymerization, is added to mouse blastocyst culture medium, hatching will not occur even when the zona pellucida is open. This inhibition is reversible once blastocysts are transferred to cytochalasin-B-free medium, indicating the importance of actin filament activity during the protrusion/extrusion processes17. As blastocysts hatch in vitro, small trophectoderm projections can be observed outside the zona pellucida. These projections exhibit ameboid movement before and after hatching is completed (Figure 7.10). Such projections have been observed in several species including mouse, human and bovine, but only at the late blastocyst stage of development16,18. Trophectoderm projections are not simple artifacts of in vitro culture; they have been noted in mouse blastocysts that have developed and hatched in vivo12,19. The length of these projections in the human averages 27 µm, and they have been suggested to serve as a first contact between blastocyst and uterus since they are localized to the precise region of the blastocyst that will ultimately attach to uterine epithelium. Blastocysts studied in vitro typically undergo repeated expansion and collapse before hatching is accomplished20,21 (Figures 7.11 and 7.12). Collapse is rapid, occurring in less than 5 min, whereas complete re-expansion requires several hours. It has been observed in our laboratory that human blastocyst collapse often occurs just before complete and final extrusion through an already ruptured zona pellucida (Figure 7.13). However, collapse of the blastocyst can be induced by mechanical, chemical, osmotic or temperature-related stress, suggesting that it is particularly susceptible to environmental

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conditions and requires careful handling. It is not known whether in vivo-produced human blastocysts collapse and re-expand in the same manner in utero. In vitro, hatching of healthy human blastocysts is typically observed on day 6 or 7 of culture, or by day 5 if the zona has been manipulated. Left in culture, blastocysts delayed in their growth may hatch as late as day 9. Because intrauterine transfer is almost always carried out by day 6 in clinical settings, it is unclear whether these are capable of implantation and subsequent normal development. In some hybrid strains of the mouse, the rate of blastocyst development rate in vitro approaches 100% and hatching rates exceed 90%, percentages far higher than the human rates of approximately 50% and 25%, respectively. Whether such differences reflect dissimilar effects of culture or different mechanisms of growth and hatching is not clear. Culture media for human in vitro fertilization almost always include protein supplements, e.g. whole serum, purified albumin or another substance which ultimately provides protein macromolecules. While considered undefined and therefore potentially toxic by some investigators, these substances have been used successfully over the years, and are held to contribute to softening and/or pliability of the zona. Conversely, culturing human preembryos in totally protein-free media has not led to widespread success, and in a number of species, including the human, adding growth factors to serum-free culture media increases blastocyst developmental and hatching rates22–24. Heparin-binding epidermal growth factor (HB-EGF) has been demonstrated to promote human blastocyst development significantly in vitro, in that blastocyst hatching rates double (45–80%), possibly through enhanced blastocyst expansion and/or by simulating preembryonic enzymes25. While human blastocysts do not express HB-EGF, it is expressed in the uterine endometrium and so may contribute to in vivo hatching success26. One must be prudent about adding these growth factors to culture media in clinical settings, however, as their effects may be harmful27. Interestingly, human blastocysts do not demonstrate remarkably higher rates of hatching when cocultured on endometrial cells, their rate of implantation after intrauterine transfer being higher than their rate of hatching in vitro28. After sperm entry into the oocyte, the zona pellucida hardens. This natural hardening helps to prevent polyspermy and serves to protect the developing preembryo. It has been speculated that the zona pellucida hardens spontaneously during in vitro culture, leading to impairment of blastocyst hatching29. To overcome this problem, various assisted hatching techniques have been developed (for review, see reference 30). Creating a slit in the zona pellucida of unfertilized oocytes by means of mechanical tearing was initially used to enhance fertilization for couples experiencing male factor infertility. However, after the creation of such a small and narrow opening, blastocysts were often observed to become trapped during subsequent hatching, leaving part inside and part outside31. This trapping phenomenon may be implicated as an occasional consequence of intracytoplasmic sperm injection (ICSI) as well (Figure 7.14). It is generally held that such an abnormal hatching process can split the inner cell mass and result in monozygotic twinning32. Therefore, acidic Tyrode’s (AT) solution has been used for assisted hatching procedures in order to create a larger opening in the zonae of day-3 preembryos. The 15– 20 µm opening this creates allows the blastocyst to hatch more freely (Figure 7.15). The consequences of AT treatment and a larger hole include: no thinning of the zona pellucida as the blastocoel enlarges, no blastocyst expansion and hatching occurring 1 day earlier than in controls. In addition, there is some evidence that these conceptuses

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implant slightly earlier than zona-intact ones33. Thus, the size of the opening and the method used to create it (mechanical vs. chemical) determine whether or not the zona pellucida thins during in vitro development and whether or not the resulting blastocyst expands (Figure 7.16). It has been our experience that preembryos undergoing assisted hatching or biopsy on day 3 demonstrate premature hatching through a thick zona on day 5 (Figure 7.17). Blastocysts whose zonae have been manipulated in these ways demonstrate extrusion wherever an artificial hole has been made, taking a path of least resistance (Figure 7.18). Initial reports of assisted hatching on day 3 followed by blastocyst transfer on day 5 demonstrate higher pregnancy and implantation rates compared to day 5 transfers with zona-intact blastocysts34. Patients with zona abnormalities, advanced maternal age, excessive cytoplasmic fragmentation or multiple previous failures have been suggested as candidates for this treatment combination35. However, these techniques often result in a smaller zona opening than after natural hatching, and therefore may result in abnormal hatching (Figure 7.19). Blastocysts cultured in vitro can demonstrate a variety of problems including the inability to hatch and excessive expansion without hatching (Figure 7.20), resulting ultimately in their collapse and degeneration (Figure 7.21). One also sees variants of partial hatching wherein blastocysts fail to complete the process and subsequently arrest and die (Figure 7.22). Occasionally, hatching occurs at more than one site, probably associated with zona damage or multiple cracks brought on by freezing and thawing procedures (Figure 7.23). To eliminate hatching problems altogether, complete removal of the zona pellucida has been attempted at various stages21. This can be accomplished through global exposure to weak AT solution or by using pronase. The AT method is rarely used because of its potential toxicity. On the other hand, pronase is widely used, the length of exposure to it depending on zona pellucida thickness. At Cornell, expanded blastocysts are exposed to 10 IU/ml of pronase (Sigma, St. Louis, MO, USA) for no more than 1 min and then washed extensively (Figure 7.24). While treatment in this manner does not appear to interfere with subsequent development or implantation, longer exposure to pronase is extremely detrimental, bringing about trophectoderm blebbing, blastocyst collapse and extensive vacuolization after 5 min36. In summary, the human hatching process in vitro involves thinning and rupture of the zona pellucida, possibly mediated by preembryonic enzymes and certainly by blastocyst expansion, followed by complete zona shedding. In vivo, uterine enzymes probably play a complementary role.

References 1. Moos J, Faundes D, Kopf GS, Schultz RM. Composition of the human zona pellucida and modifications following fertilization. Hum Reprod 1995; 10:2467–71 2. Sacco AG, Yurewicz EC, Subraminian MG, DeMayo FJ. Zona pellucida composition: species cross reactivity and contraceptive potential of antiserum to a purified pig zona antigen (PPZA). Biol Reprod 1981; 25:997–1008 3. Shabanowitz RB, O’Rand MG. Characterization of the human zona pellucida from fertilized and unfertilized eggs. J Reprod Fertil 1988; 82:151–61

An atlas of human blastocysts 214 4. Hardy K, Handyside AH, Winston RM. The human blasto cyst: cell number, death and allocation during late preimplantation development in vitro. Development 1989; 107:597–604 5. Schiewe MC, Araujo E Jr, Asch RH, Balmaceda JP. Enzymatic characterization of zona pellucida hardening in human eggs and embryos. J Assist Reprod Genet 1995; 12:2–7 6. Wright G, Wiker S, Elsner C, et al. Observations on the morphology of pronuclei and nucleoli in human zygotes and implications for cryopreservation. Hum Reprod 1990; 5:109–15 7. Chan PJ. Developmental potential of human oocytes according to zona pellucida thickness. J In Vitro Fertil Embryo Transf 1987; 4:237–41 8. Perona RM, Wassarman PM. Mouse blastocysts hatch in vitro by using a trypsin-like proteinase associated with cells of mural trophectoderm. Dev Biol 1986; 114:42–52 9. Yamazaki K, Suzuki R, Hojo E, et al. Trypsin-like hatching enzyme of the mouse blastocyst: evidence for its participation in the hatching process before zona shedding of embryos. Dev Growth Differ 1994; 36:149–54 10. Wassarman PM. Profile of a mammalian sperm receptor. Development 1990; 108:1–17 11. Schiewe MC, Hazeleger NL, Sclimenti C, Balmaceda JP. Physiological characterization of blastocyst hatching mechanisms by use of a mouse antihatching model. Fertil Steril 1995; 63:288–94 12. Lin SP, Lee RK, Tsai YJ. In vivo hatching phenomenon of mouse blastocysts during implantation. J Assist Reprod Genet 2001; 18:341–5 13. Thomas M, Jain S, Kumar GP, Laloraya M. A programmed oxyradical burst causes hatching of mouse blastocysts. J Cell Sci 1997; 110:1597–602 14. Ansell JD, Snow MH. The development of trophoblast in vitro from blastocysts containing varying amounts of inner cell mass. J Embryol Exp Morphol 1975; 33:117–85 15. Spindle Al, Pedersen RA. Hatching, attachment, and out growth of mouse blastocysts in vitro: fixed nitrogen requirements. J Exp Zool 1973; 186:305–18 16. Gonzales DS, Jones JM, Pinyopummintr T, et al. Trophectoderm projections: a potential means for locomotion, attachment and implantation of bovine, equine and human blastocysts. Hum Reprod 1996; 11:2739–45 17. Cheon YP, Gye MC, Kim CH, et al. Role of actin filaments in the hatching process of mouse blastocyst. Zygote 1999:7:123–9 18. Gonzales DS, Boatman DE, Bavister BD. Kinematics of trophectoderm projections and locomotion in the peri-implantation hamster blastocyst. Dev Dyn 1996; 205:435–44 19. McRae AC, Church RB. Cytoplasmic projections of trophectoderm distinguish implanting from preimplanting and implantation-delayed mouse blastocytes. J Reprod Fertil 1990; 88:31–40 20. Gonzales DS, Jones JM, Bavister BD, Shapiro SS. Human embryo development in vitro. Hum Reprod Update 1995; 1(item 5:video) 21. Fong CY, Bongso A, Ng SC, Kumar J, Trounson A, Ratnam S. Blastocyst transfer after enzymatic treatment of the zona pellucida: improving in-vitro fertilization and understanding implantation. Hum Reprod 1998; 13:2926–32 22. Dunglison GF, Barlow DH, Sargent IL. Leukaemia inhibitory factor significantly enhances the blastocyst formation rates of human embryos cultured in serum-free medium. Hum Reprod 1996; 11:191–6 23. Harvey MB, Kaye PL. Insulin-like growth factor-1 stimulates growth of mouse preimplantation embryos in vitro. Mol Reprod Dev 1992; 31:195–9 24. Sargent IL, Martin KL, Barlow DH. The use of recombinant growth factors to promote human embryo development in serum-free medium. Hum Reprod 1998; 13 (Suppl 4):239–48 25. Martin KL, Barlow DH, Sargent IL. Heparin-binding blastocyst development and hatching in serum-free epidermal growth factor significantly improves human medium. Hum Reprod 1998; 13:1645–52 26. Yoo HJ, Barlow DH, Mardon HJ. Temporal and spatial regulation of expression of heparinbinding epidermal growth factor-like growth factor in the human endometrium: a possible role in blastocyst implantation. Dev Genet 1997; 21:102–8

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27. Hardy K, Spanos S. Growth factor expression and function in the human and mouse preimplantation embryo. J Endocrinol 2002; 172:221–36 28. Mercader A, Simon C, Galan A, et al. An analysis of spontaneous hatching in a human endometrial epithelial coculture system: is assisted hatching justified? J Assist Reprod Genet 2001; 18:315–19 29. De Vos A, Van Steirteghem A. Zona hardening, zona drilling and assisted hatching: new achievements in assisted reproduction. Cells Tissues Organs 2000; 166:220–7 30. Zaninovic N. Assisted hatching and fragment removal. In Veeck LL, ed. An Atlas of Human Gametes and Conceptuses: an lllustrated Reference for Assisted Reproductive Technology. Carnforth, UK: Parthenon Publishing, 1999:86–96 31. Cohen J, Elsner C, Kort H, et al. Impairment of the hatching process following IVF in the human and improvement of implantation by assisting hatching using micromanipulation. Hum Reprod 1990; 5:7–13 32. Van Langendonckt A, Wyns C, Godin PA, Toussaint-Demylle D, Donnez J. Atypical hatching of a human blastocyst leading to monozygotic twinning: a case report. Fertil Steril 2000; 74:1047–50 33. Liu HC, Cohen J, Alikani M, Noyes N, Rosenwaks Z. Assisted hatching facilitates earlier implantation. Fertil Steril 1993; 60:871–5 34. Graham MC, Hoeger KM, Phipps WR. Initial IVF-ET experience with assisted hatching performed 3 days after retrieval followed by day 5 embryo transfer. Fertil Steril 2000; 74:668– 71 35. Sagoskin AW, Han T, Graham JR, Levy MJ, Stillman RJ, Tucker MJ. Healthy twin delivery after day 7 blastocyst transfer coupled with assisted hatching. Fertil Steril 2002; 77:615–17 36. Fong CY, Bongso A, Sathananthan H, Ho J, Ng SC. Ultrastructural observations of enzymatically treated human blastocysts: zona-free blastocyst transfer and rescue of blastocysts with hatching difficulties. Hum Reprod 2001; 16:540–6

Figure 7.1 Initiation of hatching in a human blastocyst on day 6. The hatching site is very near the ICM; the

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TM is of poor quality (low cell number)

Figure 7.2 Initiation of hatching in a human blastocyst on day 6. In the human, the hatching site develops in close proximity to the ICM, while in the mouse hatching occurs in an area of the mural trophectoderm, opposite to the ICM (insert)

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Figure 7.3 Hatching on day 7. The ICM is adjacent to the hatching site (ICM out of focus)

Figure 7.4 Hatching of the human blastocyst on day 6. The ICM is clearly visible at six o’clock, in close proximity to the hatching site. Onethird to one-half of the thin zona pellucida (ZP) is opened during the hatching process

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Figure 7.5 Human blastocyst showing trophectoderm vesicles (blebs) at a 1 o’clock position. These vesicles do not necessarily indicate the future hatching site and are frequently observed in ICSI-derived blastocysts (possible location of needle penetration for ICSI)

Figure 7.6 Trophectoderm vesicles in morphologically poor human blastocyst. An undeveloped ICM and

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TM can be seen. This blastocyst still possesses a somewhat thickened zona pellucida and is not yet ready to hatch. Such membrane protrusions do not indicate the onset of hatching

Figure 7.7 (a) and (b) Beginnings of the hatching process. The trophectodernn begins to herniate through a small, but true, opening (not a simple crack or vesicle breach). These blastocysts are fully expanded as indicated by their thin zona pellucidae

Figure 7.8 Human blastocyst in the process of hatching. The size of the opening is between one-third to onequarter of the circumference of the

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zona pellucida. The ICM is closely associated with the hatching site

Figure 7.9 Two sibling human blastocysts in the process of hatching. The left blastocyst exhibits a large opening in the zona pellucida and subsequently escaped easily. The one to the right possesses an already ruptured zona pellucida at a 7 o’clock position; the blastocyst itself is in a slightly contracted state just before final expansion and escape

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Figure 7.10 (a)–(f) Human blastocysts displaying small trophectoderm projections. These projections display amoeboid-like movement, suggesting they seek an implantation site. Their presence before hatching may represent the first contact between blastocyst and uterine lining. Trophectoderm projections are morphologically different to the vesicles and blebs seen in Figures 7.5 and 7.6

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Figure 7.11 (a) and (b) Human blastocysts showing partial collapse of the blastocoele. During collapse, the morphology of the ICM and TM remains unchanged. Collapse occurs frequently in in vitro cultured blastocysts, a process alternatively termed ‘blastocyst breathing’

Figure 7.12 (a) and (b) Partial collapse of the human blastocyst. The blastocyst is susceptible to environmental conditions (physical, chemical, temperature-related, or osmotic) and requires careful handling. Fragments inside the zona pellucida do

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not participate in blastocyst development

Figure 7.13 Blastocysts that collapsed, re-expanded, and subsequently hatched in vitro. (a) Collapsing blastocyst just before hatching; the zona pellucida is irregular at the hatching site (9 o’clock); (b) partial collapse before final extrusion. Note the large hatching opening; (c) blastocyst in the last stage of the hatching process. Note fragments left inside the zona pellucida; (d) hatched blastocyst. Note the increase in blastocyst size once free

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Figure 7.14 ICSI-derived blastocyst in the process of hatching. Blastocyst can become trapped due to the very small hole (<10 µm). In this instance, a typical figure-8 configuration is formed which may lead to trapping or hatching delay

Figure 7.15 High-power magnification of a hole made through an assisted hatching (AHA) procedure. (a) Side view of the opening made with acidic Tyrode’s solution; the round and very

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smooth hole was noted after blastocyst hatching. Note the large fragments remaining inside the zona pellucida; (b) frontal view of the opening. The size of the opening is between 15 and 20 µm

Figure 7.16 (a) and (b) Blastocyst hatching on day 5 after AHA was performed on day 3. No thinning of the zona peilucida occurred and blastocysts hatched earlier than usual

Figure 7.17 Blastocyst hatching on day 5 after blastomere biopsy for preimplantation genetic diagnosis on

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day 3. (a) Because an artificial hole was made, very little or no thinning of the zona pellucida occurred, resulting in the hatching of a non-expanded blastocyst; (b) in this case, the ICM was in the first section of the blastocyst to extrude through the biopsy hole

Figure 7.18 (a) and (b) Blastocyst hatching of zonae-manipulated preembryos. The blastocyst will hatch through artificially made openings in the zona pellucida, taking a course of least resistance. Zonae remain thick

Figure 7.19 (a) and (b) Premature hatching of zonae-manipulated preembryos. Due to artificial openings in the zonae, undeveloped blastocysts will hatch passively as early as late on

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day 4. Cell numbers in the TM are low and ICM formations are unclear. These undeveloped blastocysts will be exposed prematurely to the uterine environment

Figure 7.20 To the left, a very expanded human blastocyst on day 6 compared to a normally sized blastocyst to the right. The blastocyst to the left possesses an extremely thin zona pellucida without displaying any evidence of impending hatching (no rupture sites, no trophectoderm projections, not even trophectoderm vesicles). Unfortunately, these types of blastocysts often collapse and perish within their zonae, an indication for proactive zona pellucida removal

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Figure 7.21 Collapsed human blastocyst on day 7 showing degenerative changes in morphology. Inability to hatch completely resulted in blastocyst death

Figure 7.22 Human blastocyst that failed to complete hatching. Cells both outside and inside the zona pellucida show signs of impending degeneration

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Figure 7.23 Human blastocyst showing multiple blebbing sites due to cracks in the zona pellucida. This can occur after freezing and thawing procedures

Figure 7.24 (a) and (b) Human blastocysts on day 5 (a) and 6 (b) after enzymatic zona pellucida removal using pronase for 1 minute. These blastocysts are suitable for either replacement or freezing and often result in live births

8 Blastocyst cryopreservation and thawing Blastocysts have the advantage of possessing many cells. The loss of a few during freezing and thawing procedures will not compromise the integrity of the entire specimen. This may be one reason why blastocysts have been frozen and thawed so successfully over the years in domestic animals for both research and commercial purposes. Blastocyst cryopreservation in the human was first reported by Cohen and colleagues in 1985 using glycerol in a series of ten increasing concentrations1. Following that initial report, blastocyst freezing was only occasionally incorporated into clinical protocols because of the difficulties involved with maintaining high rates of blastocyst development in vitro. Eighteen years have passed since that first thawed human blastocyst led to the birth of a child, and it is 20 years since the very first pregnancy from a thawed human preembryo was reported in the world2. During this time, most IVF programs have embraced cryobiology in order to augment clinical pregnancy from a single ovarian stimulation attempt. As ovulation-induction protocols have improved, allowing the recruitment of multiple healthy oocytes, so the need has grown to manage their numbers responsibly. It is usual today to harvest in excess of ten, orsometimes even 20, mature oocytes from a woman. Before freezing techniques were routinely used in the laboratory, a woman producing so many gametes would be forced either to limit the number inseminated or to risk having to discard healthy preembryos, since only three or four could be transferred safely to the uterus after fertilization. Neither was an attractive option. It is now apparent that pregnancy after thawing is nearly equal to the transfer of fresh preembryos, at least in some programs. When the cumulative effect of adding thawed pregnancies (only from cycles failing to become pregnant following fresh transfer) to fresh pregnancies is examined, delivery outcomes are significantly enhanced3. Additionally, patients at risk of ovarian hyperstimulation syndrome (OHSS) may be managed effectively by freezing all conceptuses up front, thereby reducing, although not eliminating, the likelihood of adverse clinical symptoms if a pregnancy is established4. The availability of sequential media has led to an increase in the practice of blastocyst freezing. Several groups have reported freezing blastocysts quite successfully, some of the earliest investigations using coculture systems to support preembryo growth1,5,7. Until the mid-1990s, most reports of clinical pregnancy after thawing fell in the range of 10– 20% per transfer8, results that were not significantly better as compared to thawing earlier stages. This situation has changed dramatically, and now pregnancy rates in the range of 40–60% are common for thawed blastocysts.

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Cryobiology and the blastocyst The primary goal in establishing an appropriate freezing protocol is to do as little damage as possible while exposing blastocysts to non-physiological ultralow temperatures. Popular protocols essentially freeze-dry or dehydrate blastocysts to prevent intracellular ice from forming. The formation of intracellular ice crystals can mechanically damage specimens by disrupting and displacing organelles, or slicing through membranes. This is why freezing techniques use cryoprotective agents and control ice formation at critical temperatures. It has been shown that when human cells are placed into a medium that contains an intracellular cryoprotective agent, intracellular water readily exits the cell as a result of the higher extracellular concentration of cryoprotectant. This causes some cell shrinkage until osmotic equilibrium is reached by the slower diffusion of the cryoprotectant into the cell9. Once equilibrium is reached, the cell resumes a normal appearance. The rate of permeation of cryoprotectant and water is dependent on temperature; equilibrium is achieved faster at higher temperatures. For this reason, we have chosen to add blastocysts to cryoprotective media containing glycerol and sucrose at room temperature. However, some cryoprotectants like dimethylsulfoxide (DMSO) are toxic at elevated concentrations, and must be used at lower temperatures to reduce adverse effects. Cryoprotectants are also beneficial in their ability to lower the freezing point of a solution. Solutions may remain unfrozen at −5°C to −15°C because of supercooling (cooling to well below the freezing point without extracellular ice formation). When solutions supercool, cells do not dehydrate appropriately since there is no increase in osmotic pressure from the formation of extracellular ice crystals. To prevent supercooling, an ice crystal is introduced in a controlled fashion in a process call seeding. This contributes to intracellular dehydration as water leaves the cell to achieve equilibrium with the extracellular environment9,10. If the rate of cooling is too rapid, water cannot pass quickly enough from the cell, and as the temperature continues to drop, it reaches a point when the intracellular solute concentration is not high enough to prevent the formation of ice crystals. Blastocysts, which hold substantial intracellular water, are usually cooled at slow rates below the seeding temperature (0.3°C/min) to permit adequate dehydration. Membrane permeability by cryoprotectants varies between developmental stages. As such, it has been found that some cryoprotective agents are more suitable for blastocyst freezing than others. While DMSO and 1,2 propanediol (PROH) are frequently used for freezing early cleavage stage preembryos, propylene glycol (glycerol) is commonly used for blastocysts. All three intracellular agents have fairly small molecules that permeate cell membranes easily In addition to these, there are several extracellular substances that help dehydrate and protect cells. The most frequently used is sucrose, which possesses large, non-permeating molecules and exerts an osmotic effect to aid in accelerated cell dehydration. Sucrose cannot be used alone but is often used in conjunction with standard permeating, intracellular cryoprotectants. During the freezing procedure, all chemical reactions within cells should be suspended. Under extremely cold liquid nitrogen storage conditions (−196°C), it is estimated that it would take hundreds of years before background ionizing radiation could cause significant damage to stored cells.

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If cooling is terminated at relatively high temperatures (>−35°C), cells carry more intracellular ice than if cooled longer to lower temperatures (≤−80°C). In order to protect cells, thawing must be carried out rapidly to induce rapid ice dispersal. Conversely, samples cooled to ≤−80°C should be thawed more slowly to allow for gradual rehydration11. If water re-enters cells too rapidly, they may swell or burst. It is common to expose frozen specimens to progressively lower dilutions of cryoprotectant to ensure that it is slowly and gently removed. Vitrification The idea behind vitrification is to protect cells by completely avoiding all ice crystal formation. To accomplish this, cryoprotective solutes must be increased to 40% (wt/vol) or higher. DMSO is frequently used, but PROH, propylene glycol, ethylene glycol and other agents have been tested. Because high concentrations of these cryoprotectants are toxic at room temperature, blastocysts are generally exposed to them at 0°C. Samples may be plunged directly into liquid nitrogen without needing to introduce a seed; the viscosity is so great that solutions solidify into glasslike states. Vitrified specimens must be thawed in ice water, which is fairly inconvenient12–14. Although the procedure has been slow to gain acceptance for routine human blastocyst cryopreservation, several live births have recently been reported15–19. One novel modification to standard vitrification techniques has been to reduce the fluid content within the blastocoel before freezing19. Because the authors noted that the efficiency of their standard technique was negatively correlated to the expansion of the blastocoel, they postulated that ice crystal formation was damaging blastocysts during the cooling process. They then artificially reduced the blastocoelic volume before freezing by inserting a needle into the cavity until contraction occurred. Using this method, survival rates were much improved over those of controls, pregnancy rates showed a positive trend and implantation rates were significantly higher. Slow freezing and thawing techniques Most blastocyst freezing protocols employed today are based on the original work of Yves Menezo and co-workers, and use glycerol and sucrose as cryoprotectants20,21. The basic protocols for blastocyst freezing and thawing used by the Cornell program are illustrated in Figures 8.1 and 8.2. These are modifications of the original Menezo twostep protocols, amended in several ways to fit our current needs. Modifications include: (1) Substitution of the same base medium as is used in our phase I sequential formulation, except that it is HEPES buffered; (2) Addition of extra macromolecules (protein) in the form of 0.5 g/l human serum albumin (5% HSA solution) and 15% Plasmanate® (human plasma protein); (3) Elevation of the freezing cryoprotectant concentration to 10%; (4) Inclusion of several additional dilutions during the thawing process. Frozen-thawed blastocysts are replaced in either natural or programmed cycles. Replacement regimes are detailed in Figures 8.3–8.6. Natural cycles are not supplemented with progesterone unless there is an overwhelming reason to do so, and all

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women are treated in a prophylactic manner for 4 days with antibiotics and corticosteroids.

Blastocyst expansion and contraction Blastocysts with a high probability of survival after thaw act as perfect osmometers, shrinking, re-expanding and swelling in accordance with their osmotic environment22. One uneasy task immediately after thawing is to determine that a blastocyst has indeed survived, since it presents a contracted state for up to several hours after reincubation in standard culture medium (Figure 8.7). It has been our experience that blastocysts that shrink appropriately in response to cryoprotective agents and exhibit contracted, healthyappearing cells after thaw do quite well in their ability to survive the rigors of freezing and thawing. Zona-free blastocysts survive freezing and thawing without difficulty, indicating that the zona coat is not necessary for protective purposes during these procedures.

Pregnancy rates after blastocyst freezing and thawing Of the many tribulations associated with running a cryopreservation program, one of the most frustrating is that embryologists cannot reap the fruits of their labor (pregnancy after thawing) until months or years have passed. It is common for patients to wait for some time before returning for a thawing attempt after a negative fresh cycle, or to delay 2 or more years after the birth of a child. This situation gives rise to special problems in tracking results during a given freezing period, and makes it difficult to identify the efficiency of a new protocol. Few reports have been published to date detailing the efficiency of blastocyst freezing after culture in sequential media. Langley and colleagues describe a comparison of day-3 versus blastocyst thawed transfer during a 30-month period23. In this study involving 72 thawed blastocyst cycles, the survival rate was higher for blastocysts as compared to preembryos and the implantation rate for blastocysts was doubled (21.9% vs. 10.1%). In 2002, Behr and associates reported a 36% clinical pregnancy rate and 16% implantation rate for thawed blastocysts from 64 cycles24. Given these few peer-reviewed reports, there may not be adequate evidence to support the concept that the blastocyst stage is optimal for freezing. Nonetheless, the Cornell program has benefited greatly from the adoption of blastocyst freezing protocols. While acceptable clinical pregnancy rates in the range of 42% were realized after freezing and thawing cleavage stage preembryos in more than 800 cycles, much higher rates have been established using blastocysts (64%) without any concomitant drop in the number or proportion of patients having conceptuses frozen. Nearly one in four women under age 40 have had blastocysts frozen after undergoing day 3 transfers, and 60% of women undergoing day-5 transfers have had at least one blastocyst cryopreserved on day 5 or day 6. To date, 2000 blastocysts have been frozen. Less than one-fifth have been thawed, since so many of these patients became pregnant in their fresh cycles and have not yet returned to try for a second child.

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Most of the blastocysts frozen in the Cornell program are generated following the transfer of day-3 or day-5 conceptuses. After intrauterine transfer, remaining viable preembryos are examined each day for 1–3 additional days to evaluate their suitability for freezing. We have termed this the post-transfer observation period. Blastocysts forming on either day 5 (at least one grade 1BB) or day 6 (at least one grade 2BB) are cryopreserved for future use. Only rarely and under special circumstances have day-7 conceptuses been frozen. The survival rate for thawed blastocysts in our program is very stable at 77%. Clinical pregnancy per cycle with blastocysts thawed and replaced is 64%; the ongoing or delivered rate is 54% and the implantation rate is 39%. Pregnancy rates are not different whether blastocysts are replaced in either natural or programmed cycles. Furthermore, pregnancy rates with blastocysts are stable across all maternal ages: 14/21 women (67%) over the age of 40 have established clinical pregnancies, although their miscarriage rate is more than double that observed for younger women (24% miscarriage, 43% ongoing).

Are there differences in the reproductive potentials of blastocysts frozen on day 5 and day 6? It is generally assumed that blastocysts that develop in a timely manner in vitro are of better quality than those that develop more slowly. However, a retrospective review of blastocyst thaw outcomes from Cornell demonstrates otherwise. In our program, blastocysts have been frozen on either day 5 or day 6 depending on their speed of growth in vitro. Day-5 frozen blastocysts are thawed the day before transfer, while day-6 blastocysts are thawed in the morning when transfer is carried out. We analyzed pregnancy outcomes in 84 patients returning for thawed blastocysts over a 2-year period. Thirty-nine patients received a transfer from day-5 frozen-thawed blastocysts and 45 patients underwent transfer with day-6 blastocysts. There were no significant group differences in patient age (34.0 vs. 34.8 years, respectively), average number of blastocysts transferred (2.3 vs. 2.0) or morphology of the blastocysts after thawing. No significant differences were found in the post-thaw survival rates (73.4% vs. 80.5%), clinical pregnancy rates (63.2% vs. 63.4%) or ongoing pregnancy rates (55.3% vs. 53.7%). Nor were differences observed in implantation rates (39.8% vs. 39.5%). While it is more logical to assume that preembryos reaching the blastocyst stage faster (day 5) will be ‘healthier’ than their day-6 counterparts, these data and the data of others suggest that rate of development may not be crucial to subsequent post-thaw success24. Surprisingly, this is in direct conflict to reports of fresh transfer using day-5 and day-6 blastocysts, where pregnancy rate is observed to be significantly higher with fastergrowing conceptuses25. Also, in contrast to our work, Marek and colleagues carried out a similar study, comparing outcomes from 127 thawed blastocyst cycles where blastocysts were frozen on day 5 or day 626. Survival rates post-thawing were good for both groups, but the clinical pregnancy rate per thaw (50% vs. 29%, respectively), ongoing pregnancy rate per thaw (43% vs. 23%) and implantation rate (34% vs. 15%) were all significantly lower for day-6 blastocysts, a result quite different from what we describe.

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Children born following cryopreservation and thawing Cryopreservation has no apparent negative impact on perinatal outcome and does not appear to affect adversely the growth or health of children during infancy and early childhood27. The available data indicate that there is no elevation in the congenital malformation rate for children born after freeze-thaw procedures28–30. While it remains unclear whether freezing poses long-term risks to children so conceived or whether the freezing of blastocysts poses any additional risks over earlier stages, there is no direct evidence to raise concern.

Freezing the isolated inner cell mass It is entirely possible to isolate and freeze selectively the inner cell mass of the mouse blastocyst, a procedure that has been carried out successfully at Cornell for mouse stem cell research. In the human, an occasional blastocyst presents a healthy-appearing inner cell mass but an abnormal or degenerative trophectoderm that is clearly incapable of further development and therefore possesses no implantation potential (Figure 8.8). Whether or not the cells from these specimens are truly normal would require culture and analysis by a panel of metabolic and genetic testing procedures. Similar to this situation, an occasional thawed blastocyst will exhibit a completely degenerated trophectoderm while possessing a surviving inner cell mass (Figure 8.9). As there is no implantation potential for such blastocysts, should they not be made available for approved study? It is surely not more appropriate to discard these precious pluripotent cells rather than to grow them for potential therapeutic use.

Blastocysts before freezing and after thawing Figure 8.10 shows blastocysts with degenerative features after thaw. Figures 8.11–8.14 depict blastocysts as they appeared before freezing and again after thawing. Subsequent pregnancy outcomes are detailed.

Thawed blastocysts known to implant Figures 8.15–8.22 show thawed blastocysts that are associated with implantation success. Pregnancy outcomes are detailed.

References 1. Cohen J, Simons RF, Edwards RG, Fehilly CB, Fishel SB. Pregnancies following the frozen storage of expanding human blastocysts. J In Vitro Fertil Embryo Transf 1985; 2:59–64 2. Trounson A, Mohr L. Human pregnancy following cryopreservation, thawing and transfer of an eight-cell embryo. Nature (London) 1983; 305:707–9

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3. Veeck LL, Amundson CH, Brothman LJ, et al. Significantly enhanced pregnancy rates per cycle through cryopreservation and thaw of pronuclear stage oocytes. Fertil Steril 1993; 59:1202–7 4. Queenan JT Jr, Veeck LL, Toner JP, Oehninger S, Muasher SJ. Cryopreservation of all prezygotes in patients at risk of severe hyperstimulation does not eliminate the syndrome, but the chances of pregnancy are excellent with subsequent frozen-thaw transfers. Hum Reprod 1997; 12:1573–6 5. Hartshorne GM, Elder K, Crow J, Dyson H, Edwards RG. The influence of in-vitro development upon post-thaw survival and implantation of cryopreserved human blastocysts. Hum Reprod 1991; 6:136–41 6. Kaufman RA, Menezo Y, Hazout A, Nicollet B, DuMont M, Servy EJ. Cocultured blastocyst cryopreservation: experience of more than 500 transfer cycles. Fertil Steril 1995; 64:1125–9 7. Menezo YJ, Ben Khalifa M. Cytogenetic and cryobiology of human cocultured embryos: a 3year experience. J Assist Reprod Genet 1995:12:35–40 8. Freitas S, Le Gal F, Dzik A, et al. Value of cryopreservation of human embryos during the blastocyst stage. Contracept Fertil Sex 1994;22:396–401 9. Mazur P. Freezing of living cells: mechanisms and implications. Am J Physiol 1984;247:C125– 42 10. Whittingham DG. Some factors affecting embryo storage in laboratory animals. Ciba Found Symp 1977; 52:97–127 11. Schneider U. Cryobiological principles of embryo freezing. J In Vitro Fertil Embryo Transf 1986; 3:3–9 12. Friedler S, Shen E, Lamb EJ. Cryopreservation of mouse 2-cell embryos and ova by vitrification: methodologic studies. Fertil Steril 1987; 48:306–14 13. Friedler S, Giudice LC, Lamb EJ. Cryopreservation of embryos and ova. Fertil Steril 1988; 49:743–64 14. Quinn P, Kerin JF. Experience with the cryopreservation of human embryos using the mouse as a model to establish successful techniques. J In Vitro Fertil Embryo Transf 1986; 3:40–5 15. Choi DH, Chung HM, Lim JM, Ko JJ, Yoon TK, Cha KY. Pregnancy and delivery of healthy infants developed from vitrified blastocysts in an IVF-ET program. Fertil Steril 2000; 74:838–9 16. Yokota Y, Sato S, Yokota M, et al. Successful pregnancy following blastocyst vitrification: case report. Hum Reprod 2000; 15:1802–3 17. Yokota Y, Sato S, Yokota M, Yokota H, Araki Y Birth of a healthy baby following vitrification of human blastocysts.Fertil Steril 2001; 75:1027–9 18. Mukaida T, Nakamura S, Tomiyama T, Wada S, Kasai M, Takahashi K. Successful birth after transfer of vitrified human blastocysts with use of a cryoloop containerless technique. Fertil Steril 2001; 76:618–20 19. Vanderzwalmen P, Bertin G, Debauche C, et al. Births after vitrification at morula and blastocyst stages: effect of artificial reduction of the blastocoelic cavity before vitrification. Hum Reprod 2002; 17:744–51 20. Menezo Y, Nicollet B, Herbaut N, Andre D. Freezing cocultured human blastocysts. Fertil Steril 1992; 58:977–80 21. Menezo YJ, Nicollet B, Dumont M, Hazout A, Janny L. Factors affecting human blastocyst formation in vitro and freezing at the blastocyst stage. Acta Eur Fertil 1993; 24:207–13 22. Kaidi S, Donnay I, Lambert P, Dessy F, Massip A. Osmotic behavior of in vitro produced bovine blastocysts in cryoprotectant solutions as a potential predictive test of survival. Cryobiology 2000; 41:106–15 23. Langley MT, Marek DM, Gardner DK, Doody KM, Doody KJ. Extended embryo culture in human assisted reproduction treatments. Hum Reprod 2001; 16:902–8 24. Behr B, Gebhardt J, Lyon J, Milki AA. Factors relating to a successful cryopreserved blastocyst transfer program. Fertil Steril 2002; 77:697–9 25. Shapiro BS, Richter KS, Harris DC, Daneshmand ST. A comparison of day 5 and day 6 blastocyst transfers. Fertil Steril 2001; 75:1126–30

An atlas of human blastocysts 238 26. Marek DM, Langley MT, McKean C, Weiand L, Doody KM, Doody KJ. Frozen embryo transfer (FET) of day 5 blastocyst embryos compared to transfer of day 6 blastocyst embryos. Fertil Steril 2000; 74 (Suppl 1):S52–3 27. Wennerholm UB, Albertsson-Wikland K, Bergh C, et al. Postnatal growth and health in children born after cryopreservation as embryos. Lancet 1998; 351:1085–90 28. Wada I, Macnamee MC, Wick K, Bradfield JM, Brinsden PR. Birth characteristics and perinatal outcome of babies conceived from cryopreserved embryos. Hum Reprod 1994; 9:543– 6 29. Tarlatzis BC, Grimbizis G. Pregnancy and child outcome after assisted reproduction techniques. Hum Reprod 1999; 14(Suppl 1):231–42 30. Wennerholm WB. Cryopreservation of embryos and oocytes: obstetric outcome and health in children. Hum Reprod2000; 15(Suppl 5):18–25

Blastocyst freezing Planer biological freezer; freezing carried out in 10% glycerol/0.2 M sucrose in HEPES-based medium containing 0.5% HSA and 20% Plasmanate; use sterile 1.8 ml Nunc cryovials containing 0.3 ml freezing medium • 5% glycerol solution for 10 min • 10% glycerol/0.2 M sucrose solution for 10 min • Load into cryovials • Cool at rate of −2.0°C/min until −7.0°C • Hold 5 min, manual seed, hold 10 min • Continue cooling at −0.3°C/min until −38°C • Plunge into liquid nitrogen

Figure 8.1 Blastocyst freezing protocol used in the Cornell program Blastocyst thawing 30°C waterbath • Thaw cryovial at room temperature for 60 s • Warm cryovial in waterbath for 30–90 s (until all ice removed) • 10% glycerol+0.4 M sucrose for 30 s • 5% glycerol+0.4 M sucrose solution for 3 min • 0.4 M sucrose solution for 3 min • 0.2 M sucrose solution for 2 min • 0.1 M sucrose solution for 1 min • Wash well and incubate until transfer

Figure 8.2 Blastocyst thawing protocol used in the Cornell program

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Natural cycle replacement • Used for regular ovulatory cycles with normal luteal phase progesterone levels • Day 5 blastocysts: Thaw 4 days after LH peak or 3 days after ovulation; transfer next day • Day 6 blastocysts: Thaw 5 days after LH peak or 494 days after ovulation; transfer same day • No supplemental progesterone unless indicated or after previous failure without supplementation • Begin administering medrol and tetracycline on the day of the LH surge; continue for 4 days

Figure 8.3 Cornell replacement strategy for natural cycle thawed blastocyst transfer Natural cycle replacement Progesterone (if indicated): 200 mg micronized P4 vaginally b.i.d. or t.i.d.; continued until negative pregnancy test 14 days after replacement or through week 12 if pregnant (weaned down starting weeks 9–10) Medrol: 16 mg/day for 4 days starting day of LH surge Tetracycline: 250 mg q.i.d. for 4 days starting day of LH surge

Figure 8.4 Cornell replacement regimen for natural cycle thawed blastocyst transfer Programmed replacement (adequate suppression confirmed on day 2 of cycle) • Luteal suppression with 0.2 mg GnRHa; drop to 0.1 mg starting on predetermined day 1 and maintain until day 15 • Transdermal estrogen patches:

Days 1–4 0.1 mg every other day Days 5–8 0.2 mg every other day Days 9–14 0.3–0.4 mg every other day (depending on E2 levels)

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Days 15+0.2 mg (every other day until negative pregnancy test or for 7 weeks if pregnant) • 50 mg P4 day 15 through 12 weeks gestation (weaned down starting week 9–10, depending on serum levels) • Tetracycline+Medrol beginning day 15 for 494 days

Day 5 blastocysts: Thaw day 19, transfer next day Day 6 blastocysts: Thaw day 20, transfer same day Figure 8.5 Cornell replacement strategy for programmed cycle thawed blastocyst transfer Programmed replacement Estrogen patches: Climara, 0.1 mg patch (every other day) Progesterone: 50mg/day i.m. beginning day 15; continued until negative pregnancy test or through week 12 if pregnant (weaned starting weeks 9–10) Medrol: 16 mg/day for 4 days starting day 15 Tetracycline: 250 mg q.i.d. for 4 days starting day 15

Figure 8.6 Cornell replacement regime for programmed cycle thawed blastocyst transfer

Figure 8.7 Thawed blastocysts. (a) Contracted blastocyst immediately after thawing; (b) contracted, hatched blastocyst 30 minutes after thawing

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Figure 8.8 Non-surviving blastocyst exhibiting a degenerative trophectoderm immediately after thawing; inner cell mass appears viable

Figure 8.9 Viable and proliferating inner cell mass surrounded by a degenerating trophectoderm after thawing; the process of hatching is not completed and blastocyst appears trapped

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Figure 8.10 Degenerative features after thawing. (a) Contracted cavitating morula with degenerating fragments after thaw: (b) non-viable inner cell mass despite blastocoel re-expansion after thaw

Figure 8.11 Before freezing and after thawing. Maternal age 39 years at time of freeze. (a) Expanded blastocyst immediately before freezing; (b) same blastocyst as in (a) after thawing 141 days later; membrane blebbing through the zona pellucida is seen, likely by virtue of subtle zona pellucida damage that occurred during thaw. This blastocyst was transferred, implanted, and a pregnancy with one fetal heart is on-going >30 weeks

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Figure 8.12 Before freezing and after thawing. Maternal age 31 years at time of freeze. (a) Expanded blastocyst immediately before freezing; (b) same blastocyst as in (a) after thawing 163 days later; inner cell mass appears intact but some cells of the trophectoderm appear unhealthy. This blastocyst was transferred, implanted, but an early loss occurred after ultrasound evidence of one gestational sac

Figure 8.13 Before freezing and after thawing. Maternal age 22 years at time of freeze. (a) Expanded blastocyst immediately before freezing; proliferative trophectoderm and smallish inner cell mass; (b) same blastocyst as in (a) after thawing 124

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days later; most cells appear viable. This blastocyst was transferred, implanted, and a healthy male child was delivered at term

Figure 8.14 Before freezing and after thawing. Maternal age 37 years at time of freeze. (a) Expanded blastocyst immediately before freezing; inner cell mass is large and trophectoderm is sparse; (b) same blastocyst as in (a) after thawing 56 days later; most cells appear viable. This blastocyst was transferred, implanted, and a healthy female child was delivered. Note here that the blastocyst was thawed almost 1 full day before replacement and that the trophectoderm appears quite different after prolonged culture, being made up of many more cells

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Figure 8.15 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 57 days, this blastocyst was thawed and led to the birth of a healthy female child. Maternal age 34 years at time of freeze

Figure 8.16 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 79 days, this blastocyst was thawed and transferred. A currently ongoing pregnancy was established with one fetal heart. Maternal age 26 years at time of freeze

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Figure 8.17 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 155 days, this blastocyst was thawed and transferred. Acurrently ongoing pregnancy was established with one fetal heart. Maternal age 41 years at time of freeze

Figure 8.18 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 558 days, this blastocyst was thawed and transferred. A pregnancy was established with one fetal heart. One healthy male was delivered. Maternal age 30 years at time of freeze

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Figure 8.19 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 102 days, this blastocyst was thawed and transferred. A pregnancy was established with one fetal heart. One healthy female was delivered. Maternal age 34 years at time of freeze

Figure 8.20 Blastocyst known to have implanted after freeze, thaw, and transfer. After cryostorage for 86 days, this blastocyst was thawed and transferred. A pregnancy was established with one fetal heart. One healthy male was delivered. Maternal age 39 years at time of freeze

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Figure 8.21 Two blastocysts known to have implanted after freeze, thaw, and transfer. After cryostorage for 84 days, these blastocysts were thawed and transferred. Two healthy male children were subsequently delivered. Maternal age 29 years at time of freeze

Figure 8.22 Two blastocysts known to have implanted after freeze, thaw, and transfer. After cryostorage for 68 days, these blastocysts were thawed and transferred. Two fetal hearts have been documented for this currently ongoing pregnancy. Maternal age 37 years at time of freeze

9 Cell death (apoptosis) in human blastocysts Kate Hardy, Sophie Spanos and David L.Becker

INTRODUCTION The orchestrated death of cells is fundamental for tissue and organ modelling during development1–3. Furthermore, the ability of cells to die without damaging adjacent cells is a vital component of the quality control and repair programs needed for maintaining healthy tissue. In this way, excess, damaged or unwanted cells can be safely and discreetly removed. These key concepts of developmental biology originated from a seminal article published in 1972 by Kerr and colleagues4, who first observed and described the classic morphological features of this physiological mode of cell death, which they termed apoptosis. Examples of developmental events that require extensive apoptosis include the sculpting of digits in the developing limb bud, and the removal of the Müllerian duct during development of the male3. During early rodent development, apoptosis plays a crucial role in amniotic cavity formation, soon after implantation5. Surprisingly, there is now increasing evidence that apoptosis occurs before implantation (for review, see reference 6). Apoptotic nuclei have been seen in blastocysts from many mammalian species, including mouse7–9, rat10, cow11–13, baboon14, rhesus monkey15–17 and human11,18–22. These dying cells are characterized by blebbing of the nuclear membrane, chromatin condensation, cytoplasmic vacuoles and the formation of both nuclear and cytoplasmic fragments.

APOPTOSIS Morphological features of apoptosis Cells can die either as a result of accidental injury or by design. Injury, such as physical trauma or ischemia, will result in the death of swathes of cells, which will undergo swelling and membrane rupture2. The cellular contents will spill into the extracellular spaces, invoking an inflammatory response in adjacent healthy tissue. This process is known as necrosis. However, cells also have mechanisms in place which allow them to control and contain their death, and to die without causing widespread chaos. Death by this mechanism is sufficiently neat and quick to be almost undetectable by classical histology, and it was only 30 years ago that the extent of this phenomenon, which came to be termed apoptosis, was recognized and described4. Apoptosis is the mechanism by which cells undergo a programmed and carefully orchestrated series of steps whereby the

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cellular contents are dismantled, packaged and disposed of with no associated inflammation. Apoptosis is characterized by a sequence of morphologically distinct phases, which distinguish it from necrosis (Figure 9.1). Apoptosis is heralded by cell rounding, indicating that changes in cell adhesion occur2. Initially chromatin condenses into clumps on the inner nuclear membrane and the cytoplasm also condenses. The nuclear and cytoplasmic membranes become indented and the nucleus fragments. Finally, the whole cell blebs and fragments into membrane-bound apoptotic bodies, which may contain nuclear fragments. The production of membrane-bound apoptotic bodies prevents the release of intracellular contents from the dying cell, which would result in damage to adjacent healthy cells and inflammation. These bodies are either dispersed in the intercellular tissue spaces and extruded from the tissue or phagocytosed by neighboring tissue cells. Apoptotic cells that are not phagocytosed, for example those shed from an epithelium into a duct lumen, undergo secondary necrosis. This classical sequence of events has been seen at the ultrastructural level in a wide variety of different cell types2,23,24, and is under physiological control. Regulation and execution of apoptosis Apoptosis may result from activation of an endogenous developmental program, or be induced by a stimulus, such as DNA damage, cell surface ligand-receptor interactions, disruption of cell-cell or cell-matrix interactions, or growth factor deprivation. The signals that initiate apoptosis mediate their effects through distinct signal transduction pathways which trigger the apoptotic machinery via a variety of routes, but which all have the same end point: the death of the cell. The co-ordinated dismantling of a cell by apoptosis involves a large family of death proteases known as the caspases. All caspase proteins exist normally in healthy cells as inactive proenzymes, which require proteolysis for enzymatic activation. Activation of these caspases results in the death of the cell. Caspases can be broadly subdivided into two groups: initiator caspases (caspase 8 and caspase 9), whose main function is to activate downstream caspases, and effector caspases (such as caspases 3, 6 and 7), which are responsible for dismantling cellular proteins critical for cell survival, including those involved in the cytoskeleton, DNA repair, nuclear envelope integrity and cell cycle control23. The activation of effector caspases results in the apoptotic morphology observed in cells, including the formation of fragmented nuclei and apoptotic bodies25. Effector caspases also activate endonucleases, resulting in the cleavage of DNA between nucleosomes into oligonucleosomal fragments2,26,27. The other major regulators of apoptosis are the bcl-2 family of mitochondrial proteins, various members of which promote (bad, bax, bak, bcl-xs) and inhibit (bcl-2, bcl-w, bclxL) apoptosis. It is thought that the ratio between pro- and antiapoptotic members determines whether the cell lives or dies28. The bcl-2 family is believed to regulate the release of proapoptotic factors from mitochondria, which in turn leads to the activation of caspases29.

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Evidence for apoptosis in preimplantation embryos The study of apoptosis in preimplantation human embryos is constrained by the limited number of embryos available for research, and in most studies the only embryos available are those which have arrested development or which are highly fragmented and unsuitable for cryopreservation. Furthermore, each embryo contains only between one and 100 or so cells. It is difficult to estimate the full extent of cell death in the mammalian embryo, as it is not known how long dead cells persist; therefore, current techniques provide only a ‘snapshot’ of the embryo at a specific time. Historically, while it was evident from histological analysis that cell death was taking place in mammalian blastocysts, it was not clear whether the death was necrotic or apoptotic. As around 50% of human embryos fail to reach the blastocyst stage22, and as the involvement of cell death in this developmental arrest is a possibility30, the importance of understanding the causes and roles of these processes in the human embryo cannot be underestimated. Membrane changes Early apoptotic events involve changes in the cell membrane, which are controlled by initiator caspases. These include modifications of the plasma membrane itself, and alterations in the cytoskeleton leading to aberrant membrane behavior such as blebbing (reviewed in reference 31). An early event of apoptosis is a redistribution of the phospholipid phosphatidylserine in the cell membrane. In healthy cells, phosphatidylserine is normally confined to the inner cytoplasmic leaflet of the membrane. In apoptotic cells this asymmetry is lost, with exposure of phosphatidylserine on the outer leaflet32. Annexin V (an anticoagulant protein) has a specific and high affinity for phosphatidylserine, and labelled annexin V is a classical marker for externalized phosphatidylserine, indicative of the early stages of apoptosis. Annexin V labelling has been examined in early human embryos and oocytes33–35, with conflicting results. Levy and colleagues33 found extensive labelling in arrested and fragmented human embryos. However, the nuclei in these embryos also labelled with propidium iodide, indicating that the plasma membrane was permeable. Therefore, it was probable that these arrested embryos were undergoing secondary necrosis. In contrast, Antczak and Van Blerkom35 found no annexin V or propidium iodide labelling in cleavage-stage human embryos with varying degrees of fragmentation, strongly suggesting that there is no correlation between fragmentation and apoptosis. Video time-lapse cinematography is a useful tool for observing the kinetics of apoptosis. Using this technique, violent membrane blebbing in trophectoderm and inner cell mass cells over a time period of up to 1 h, followed by rupture of the cell and its contents into the blastocoel cavity, have been observed (Figure 9.2). Similar cytoplasmic blebbing has been seen in other cell types undergoing apoptosis using time-lapse cinematography23,36. In addition, small cytoplasmic blebs have been seen on the surface of the inner cell mass (e.g. Figure 9.3), which may either reflect these early membrane events, or be the final products of apoptosis in these cells.

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Chromatin condensation The earliest morphological change indicative of apoptosis, which in fact is a late event within the apoptosis cascade, is the condensation of chromatin on the inner nuclear membrane. Initially, this was detected by transmission electron microscopy in mouse embryos7 and later in human embryos20,37. More recently, nuclei with clumped or condensed chromatin have been visualized in arrested cleavage stage embryos33,38 and human blastocysts using propidium iodide or Hoescht staining and standard fluorescence microscopy19 or laser scanning confocal microscopy39 (Figures 9.4–9.8). Condensed nuclei With the same approach of labelling DNA, condensed and misshapen nuclei have been seen in fragmented arrested20,33 and developing21 human embryos. These may be precursors of fragmented nuclei (see below). Nuclear fragmentation Nuclear fragmentation, which is perhaps the most classical morphological feature of apoptosis, has been observed in human embryos using a variety of techniques. Fragmented nuclei have been described in paraffin-embedded sections of human embryos in vivo40, providing unique evidence that apoptosis occurs in vivo during preimplantation human development. Fragmenting nuclei have also been visualized by fluorescence microscopy of blastocysts labelled with DNA-specific fluorochromes19. Apoptotic nuclei in both the trophectoderm and the inner cell mass can be identified as discrete clusters of fluorochrome-labelled nuclear fragments, which are smaller than intact healthy nuclei (Figures 9.7 and 9.8). More recently, confocal microscopy has allowed clear identification and quantification of fragmented nuclei at high resolution in human blastocysts21,22 (Figures 9.9–9.11). DNA fragmentation Another classical feature of apoptosis is the degradation of DNA into oligonucleosomal fragments. The small numbers of cells in preimplantation embryos do not allow the use of electrophoretic techniques to look for DNA laddering typical of apoptotic nuclei. However, the development of terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling (TUNEL)41 allows the assessment of nuclear DNA fragmentation in situ. This technique can be used on specimens with only a few cells, and is based on the fluorescent labelling of the 3' end of oligonucleosome fragments. This technique has the additional advantages of allowing both the localization and the quantification of the percentage of nuclei with DNA fragmentation41. The enzyme terminal deoxynucleotidyl transferase (TdT) binds to exposed 3'-OH ends of DNA single-strand breaks and catalyzes the addition of labelled deoxynucleotides that have been labelled with a marker such as fluorescein isothiocyanate (FITC), in which case TUNEL-positive nuclei will be clearly seen using fluorescent microscopy or laser scanning confocal microscopy (Figures 9.10–9.17).

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Positive labelling of fragmented nuclei has been observed in arrested20,33 and fragmented42 cleavage-stage human embryos, and in human blastocysts21,22, providing evidence that DNA is being degraded into oligonucleosomal fragments. Where TUNEL labelling has been seen in arrested embryos, it is possible that DNA degradation is occurring in cells undergoing the early stages of secondary necrosis, rather than apoptosis. Certainly secondary necrotic changes with disrupted membranes and swollen organelles have been observed in cytoplasmic fragments37. During ‘normal’ development of non-arrested embryos in vitro, DNA fragmentation is only seen at the blastocyst stage in mice43, after compaction in human embryos22 and after the 8-cell stage in cow embryos13. Condensed cytoplasm Condensed cytoplasm has been observed by transmission electron microscopy in arrested embryos20,37,42. Changes in expression of genes involved in regulation and execution of apoptosis The classical morphological changes associated with apoptosis, including nuclear fragmentation, are a consequence of the action of active caspases, which in turn are regulated, at least in part, by the bcl-2 family of proteins. Expression of mRNA and protein for members of the caspase and bcl-2 families of genes in preimplantation embryos have been studied using reverse transcriptasepolymerase chain reaction (RT-PCR) and immunohisto-chemistry, providing information about the timing of gene expression. Studies in mouse preimplantation embryos have shown that preimplantation oocytes and embryos of all stages express the relevant molecular machinery to undergo apoptosis44,45. In addition to members of the bcl-2 family, caspase expression has been detected in mouse embryos throughout preimplantation development45. Quantitative RT-PCR has been used to detect increased expression of bax mRNA in mouse embryos exposed to high glucose concentrations in the culture medium46. In human preimplantation embryos, mRNA for bax, bcl-2 and bad has also been detected by RT-PCR47,48. In addition, bax, bcl-2, bcl-x and bcl-w protein has been detected in human embryos using immunohisto-chemistry35,39,48,49 (Figures 9.18 and 9.19). The presence of active caspases within cells indicates that the execution phase of the apoptotic cascade is under way. Detection of active caspases currently involves the use of fluorescently tagged specific caspase inhibitors that bind only to active caspases. The cytoplasm of cells which contain active caspases, and are undergoing apoptosis, is fluorescently labelled. Healthy cells remain unlabelled. Using this approach, active caspases have not been detected in arrested or developing cleavage-stage human embryos, but have been seen in some (but not all) fragments48,50. However, following compaction, individual cells with cytoplasmic labelling for active caspases have been seen in morulae and blastocysts48 (Figures 9.20 and 9.21). In particular, cells with

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fragmented nuclei are frequently positive for active caspases, suggesting that they play a role in the formation of the apoptotic morphology seen in human blastocysts. Morphological changes Excluded cells The earliest events in apoptosis involve the cell membrane, with the apoptotic cell rounding up and separating from neighboring cells23. At the time of compaction and blastocyst formation, it is not uncommon to see excluded cells which lie between the developing embryo and the zona pellucida (Figures 9.22 and 9.23) or within the blastocoel cavity (Figure 9.24). Transmission electron microscopy and laser scanning confocal microscopy have often revealed extruded cells in the perivitelline space or blastocoel cavity12,18. Cytoplasmic fragmentation Over 75% of human embryos generated following in vitro fertilization (IVF) have various degrees of cytoplasmic fragmentation (Figure 9.25). Fragmentation has also been seen in human embryos in vivo40,51,52, indicating that fragmentation is not an in vitro artifact. Ultrastructural examination of fragmented embryos in vitro has shown that these membrane-bound fragments contain organelles and resemble apoptotic bodies20,42. The issue of whether cytoplasmic fragments are apoptotic bodies or not remains contentious, with no clear consensus. Some workers have shown that features of apoptosis are present in fragments; either fragmented DNA20, active caspases48,50 or ultrastructural features20. In contrast, others have failed to 5demonstrate either TUNEL labelling (indicating DNA fragmentation) or annexin V labelling (indicating phosphatidylserine redistribution in the plasma membrane)35, leading these workers to postulate that fragmentation was not part of, or a consequence of, apoptosis. It is not known how long fragments have been present, and whether they arose during the previous cleavage division or during earlier ones. If the fragments have persisted for some time it is possible that the DNA has fragmented during secondary necrosis. Furthermore, fragmentation is not inhibited by caspase inhibitors53, providing further evidence that the mechanisms involved in fragment formation do not invariably involve apoptosis. Cytoplasmic vacuoles Translucent vacuoles are frequently associated with cytoplasmic condensation typical of cells undergoing apoptosis2. Vacuoles are a common feature of human preimplantation embryos and have been seen by light38 (Figures 9.4 and 9.26), electron7,54 and laser scanning confocal38 microscopy.

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Cell corpses Cell corpses have been observed by transmission electron microscopy, lying between cells11,37. Phagocytosis Further degeneration results in the formation of extracellular debris2, with the presence of mitochondria and nuclear remnants confirming its cellular origin, for example, in arrested human embryos20. Confocal microscopy of human and cow blastocysts has shown cells being engulfed by neighboring cells38,55 (Figure 9.27). Autophagic vacuoles have been seen at the ultrastructural level using transmission electron microscopy in human11, cow55, rhesus16 and mouse blastocysts7. However, while it appears that many dead cells are cleared by phagocytosis within the blastocyst, other apparently arrested cells simply persist. In other cell types, apoptotic cells formed in single-layered epithelia may be extruded into lumina, andthus escape phagocytosis2,24. This process appears to occur in human blastocysts, with isolated cells being observed in the blastocoel cavity or between the trophectoderm and the zona pellucida18,39 (Figures 9.22–9.24). These cells and fragments are excluded from normal development. These isolated cells in the blastocoel cavity have been shown to be TUNEL labelled39, as have similar cells in mouse blastocysts43. In some cases, these cells are large, and this, coupled with the lack of intercellular junctions, poorly differentiated mitochondria and paucity of rough endoplasmic reticulum, suggests that these cells originated during early cleavage. It is possible that such cells lack cell surface markers which would promote their ingestion by neighboring cells, causing them to persist throughout preimplantation development. Furthermore, cell corpses with condensed chromatin and cytoplasm were not phagocytosed37. Timing of apoptosis In human embryos which are non-arrested and developing on schedule, apoptosis (as quantified by DNA and nuclear fragmentation) has not been seen in early cleavage stages22,42. Nuclear fragmentation was observed rarely before compaction and at increasing levels during blastocyst formation. TUNEL labelling was seen only after compaction, and the proportion of nuclei with DNA fragmentation increased during blastocyst formation to around 10%22. This is consistent with studies of rodent, bovine and porcine embryos, where apoptosis is not observed until the morula and blastocyst stages9,10,12,43,56. It has been proposed that upstream death inducers or downstream death effectors are either absent or suppressed during early cleavage stages12. As it has been shown that most components of the apoptotic machinery are present in mouse44,45 and human48 embryos throughout development, the most likely scenario is that apoptosis is suppressed during early cleavage stages.

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Incidence of apoptosis In the human blastocyst, cell death is present equally in the trophectoderm and the inner cell mass, with approximately 10% of nuclei in each lineage showing evidence of apoptosis. The incidence appears to be correlated with embryo quality, with embryos of poor morphology having higher levels of apoptosis19. Role of apoptosis In addition to cell division and differentiation within the preimplantation embryo, it is now clear that cell death by apoptosis is common, even in vivo9. Within a blastocyst, the percentage of dying cells is sufficiently high (10–20%) to suggest that apoptosis is playing a developmental role, which has not yet been elucidated. It is possible that the higher levels of apoptosis seen in the inner cell mass of the mouse blastocyst are to remove cells that retain the potential to form trophectoderm57, which could lead to ectopic expression of trophectoderm cells in the germ layers during later development. In addition, levels of apoptosis can be modulated by environmental factors. Apoptosis in preimplantation embryos is increased in suboptimal culture media58, in the presence of high concentrations of glucose46,59 and in the absence of specific growth factors21,43,60,61. Furthermore, apoptosis has been associated with increased levels of reactive oxygen species thought to result from high oxygen tension during culture42, exposure to high concentrations of sperm during human IVF and increased maternal age62. Finally, nuclear63 and chromosomal64,65 abnormalities are common in human embryos, and it has been suggested that one role of apoptosis may be to remove such defective cells6. In conclusion, apoptosis occurs in human embryos following compaction. The reasons for certain cells undergoing apoptosis are unclear, the role during preimplantation development remains unknown and the early signalling events that trigger apoptosis require elucidation.

References 1. Glücksmann A. Cell death in normal development. Arch Biol 1965; 76:419–37 2. Wyllie AH, Kerr JF, Currie AR. Cell death: the significance of apoptosis. Int Rev Cytol 1980; 68:251–306 3. Meier P, Finch A, Evan G. Apoptosis in development. Nature (London) 2000; 407:796–801 4. Kerr JF, Wyllie AH, Currie AR. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer 1972; 26:239–57 5. Coucouvanis E, Martin GR. Signals for death and survival: a two-step mechanism for cavitation in the vertebrate embryo. Cell 1995; 83:279–87 6. Hardy K. Cell death in the mammalian blastocyst. Mol Hum Reprod 1997; 3:919–25 7. El-Shershaby AM, Hinchliffe JR. Cell redundancy in the zona-intact preimplantation mouse blastocyst: a light and electron microscope study of dead cells and their fate. J Embryol Exp Morphol 1974; 31:643–54 8. Copp AJ. Interaction between inner cell mass and trophectoderm of the mouse blastocyst. I. A study of cellular proliferation. J Embryol Exp Morphol 1978; 48:109–25 9. Handyside AH, Hunter S. Cell division and death in the mouse blastocyst before implantation. Roux’s Arch Dev Biol1986;195:519–26

An atlas of human blastocysts 258 10. Pampfer S, De Hertogh R, Vanderheyden I, Michiels B, Vercheval M. Cell allocation to the inner cell mass and the trophectoderm in rat embryos during in vivo preimplantation development. Roux’s Arch Dev Biol 1990; 198:257–63 11. Mohr LR, Trounson AO. Comparative ultrastructure of hatched human, mouse and bovine blastocysts. J Reprod Fertil 1982; 66:499–504 12. Matwee C, Betts DH, King WA. Apoptosis in the early bovine embryo. Zygote 2000; 8:57–68 13. Byrne AT, Southgate J, Brison DR, Leese HJ. Analysis of apoptosis in the preimplantation bovine embryo using TUNEL. J Reprod Fertil 1999; 117:97–105 14. Enders AC, Lantz KC, Schlafke S. Differentiation of the inner cell mass of the baboon blastocyst. Anat Rec 1990; 226:237–48 15. Hurst PR, Jefferies K, Eckstein P, Wheeler AG. An ultra structural study of preimplantation uterine embryos of the rhesus monkey. J Anat 1978; 126:209–20 16. Enders AC, Schlafke S. Differentiation of the blastocyst of the rhesus monkey. Am J Anat 1981; 162:1–21 17. Enders AC, Hendrickx AG, Binkerd PE. Abnormal development of blastocysts and blastomeres in the rhesus monkey. Biol Reprod 1982; 26:353–66 18. Lopata A, Kohlman DJ, Kellow GN. The fine structure of human blastocysts developed in culture. In Embryonic Development, Part B: Cellular Aspects. New York: Alan R Liss, 1982:69–85 19. Hardy K, Handyside AH, Winston RM. The human blastocyst: cell number, death and allocation during late preimplantation development in vitro. Development 1989; 107:597–604 20. Jurisicova A, Varmuza S, Casper RF. Programmed cell death and human embryo fragmentation. Mol Hum Reprod 1996; 2:93–8 21. Spanos S, Becker DL, Winston RM, Hardy K. Antiapoptotic action of insulin-like growth factor-l during human preimplantation embryo development. Biol Reprod 2000; 63:1413–20 22. Hardy K, Spanos S, Becker D, lannelli P, Winston RM, Stark J. From cell death to embryo arrest: mathematical models of human preimplantation embryo development. Proc Natl Acad Sci USA 2001; 98:1655–60 23. Wyllie AH. Apoptosis: an overview. Br Med Bull 1997; 53:451–65 24. Harmon BV, Winterford CM, O’Brien BA, Allan DJ. Morphological criteria for identifying apoptosis. In Celis JE, ed. Cell Biology: A Laboratory Handbook, 2nd edn. San Diego: Academic Press, 1998; 1:327–40 25. Earnshaw WC, Martins LM, Kaufmann SH. Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu Rev Biochem 1999; 68:383–424 26. Nagata S. Apoptotic DNA fragmentation. Exp Cell Res 2000; 256:12–18 27. Arends MJ, Wyllie AH. Apoptosis: mechanisms and roles in pathology. Int Rev Exp Pathol 1991; 32:223–54 28. Oltvai ZN, Milliman CL, Korsmeyer SJ. Bcl-2 heterodimerizes in vivo with a conserved homolog, Bax, that accelerates programmed cell death. Cell 1993; 74:609–19 29. Adams JM, Cory S. Life-or-death decisions by the Bcl-2 protein family. Trends Biochem Sci 2001; 26:61–6 30. Jurisicova A, Varmuza S, Casper RF. Involvement of programmed cell death in preimplantation embryo demise. Hum Reprod Update 1995; 1:558–66 31. Huppertz B, Frank HG, Kaufmann P. The apoptosis cascade—morphological and immunohistochemical methods for its visualization. Anat Embryol (Berl) 1999; 200:1–18 32. Martin SJ, Reutelingsperger CP, McGahon AJ, et al. Early redistribution of plasma membrane phosphatidylserine is a general feature of apoptosis regardless of the initiating stimulus: inhibition by overexpression of Bcl-2 and Abl. J Exp Med 1995; 182:1545–56 33. Levy R, Benchaib M, Cordonier H, Souchier C, Guerin JF. Annexin V labelling and terminal transferase-mediated DNA end labelling (TUNEL) assay in human arrested embryos. Mol Hum Reprod 1998; 4:775–83

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34. Van Blerkom J, Davis PW. DNA strand breaks and phosphatidylserine redistribution in newly ovulated and cultured mouse and human oocytes: occurrence and relationship to apoptosis. Hum Reprod 1998; 13:1317–24 35. Antczak M, Van Blerkom J. Temporal and spatial aspects of fragmentation in early human embryos: possible effects on developmental competence and association with the differential elimination of regulatory proteins from polarized domains. Hum Reprod 1999; 14:429–47 36. Collins JA, Schandi CA, Young KK, Vesely J, Willingham MC. Major DNA fragmentation is a late event in apoptosis. J Histochem Cytochem 1997; 45:923–34 37. Jurisicova A, Varmuza SL, Casper RF. Developmental consequences of programmed cell death in human preimplantation embryos. In Tilly J, ed. Cell Death in Reproductive Physiology. New York: Springer Verlag, 1997:34–47 38. Hardy K, Warner A, Winston RM, Becker DL. Expression of intercellular junctions during preimplantation development of the human embryo. Mol Hum Reprod 1996; 2:621–32 39. Hardy K. Apoptosis in the human embryo. Rev Reprod 1999; 4:125–34 40. Hertig AT, Rock J, Adams EC, Mulligan WJ. On the preimplantation stages of the human ovum: a description of four normal and four abnormal specimens ranging from the second to fifth day of development. Contrib Embryol 1954; 35:199–220 41. Gavrieli Y, Sherman Y, Ben-Sasson SA. Identification of programmed cell death in situ via specific labeling of nuclear DNA fragmentation. J Cell Biol 1992; 119:493–501 42. Yang HW, Hwang KJ, Kwon HC, Kim HS, Choi KW, Oh KS. Detection of reactive oxygen species (ROS) and apoptosis in human fragmented embryos. Hum Reprod 1998; 13:998–1002 43. Brison DR, Schultz RM. Apoptosis during mouse blastocyst formation: evidence for a role for survival factors including transforming growth factor α. Biol Reprod 1997; 56:1088–96 44. Jurisicova A, Latham KE, Casper RF, Casper RF, Varmuza SL. Expression and regulation of genes associated with cell death during murine preimplantation embryo development. Mol Reprod Dev 1998; 51:243–53 45. Exley GE, Tang C, McElhinny AS, Warner CM. Expression of caspase and BCL-2 apoptotic family members in mouse preimplantation embryos. Biol Reprod 1999; 61:231–9 46. Moley KH, Chi MM, Knudson CM, Korsmeyer SJ, Mueckler MM. Hyperglycemia induces apoptosis in pre-implantation embryos through cell death effector pathways. Nat Med 1998; 4:1421–4 47. Warner CM, Cao W, Exley GE, et al. Genetic regulation of egg and embryo survival. Hum Reprod 1998; 13 (Suppl 3):178–90, discussion 191–6 48. Spanos S, Rice S, Karagiannis P, et al. Caspase activity and expression of cell death genes during development of human preimplantation embryos. Reproduction 2002; 124:353–63 49. Brison DR. Apoptosis in mammalian preimplantation embryos: regulation by survival factors. Hum Fertil 2000; 3:36–47 50. Martinez F, Rienzi L, lacobelli M, et al. Caspase activity in preimplantation human embryos is not associated with apoptosis. Hum Reprod 2002; 17:1584–90 51. Pereda J, Croxatto HB. Ultrastructure of a seven-cell human embryo. Biol Reprod 1978; 18:481–9 52. Buster JE, Bustillo M, Rodi IA, et al. Biologic and morphologic development of donated human ova recovered by nonsurgical uterine lavage. Am J Obstet Gynecol 1985; 153:211–17 53. Xu J, Cheung T, Chan ST, Ho P, Yeung WS. The incidence of cytoplasmic fragmentation in mouse embryos in vitro is not affected by inhibition of caspase activity. Fertil Steril 2001; 75:986–91 54. Sathananthan AH, Wood C, Leeton JF. Ultrastructural evaluation of 8–16 cell human embryos cultured in vitro. Micron 1982; 13:193–203 55. Plante L, King WA. Light and electron microscopic analysis of bovine embryos derived by in vitro and in vivo fertilization. J Assist Reprod Genet 1994; 11:515–29

An atlas of human blastocysts 260 56. Long CR, Dobrinsky JR, Garrett WM, Johnson LA. Dual labeling of the cytoskeleton and DNA strand breaks in porcine embryos produced in vivo and in vitro. Mol Reprod Dev 1998; 51:59– 65 57. Pierce GB, Lewellyn AL, Parchment RE. Mechanism of programmed cell death in the blastocyst. Proc Natl Acad Sci USA 1989; 86:3654–8 58. Devreker F, Hardy K. Effects of glutamine and taurine on preimplantation development and cleavage of mouse embryos in vitro. Biol Reprod 1997; 57:921–8 59. Pampfer S, Vanderheyden I, McCracken JE, Vesela J, De Hertogh R. Increased cell death in rat blastocysts exposed to maternal diabetes in utero and to high glucose or tumor necrosis factor-α in vitro. Development 1997; 124:4827–36 60. Herrler A, Krusche CA, Beier HM. Insulin and insulin-like growth factor-l promote rabbit blastocyst development and prevent apoptosis. Biol Reprod 1998; 59:1302–10 61. O’Neill C. Autocrine mediators are required to act on the embryo by the 2-cell stage to promote normal development and survival of mouse preimplantation embryos in vitro. Biol Reprod 1998; 58:1303–9 62. Jurisicova A, Rogers I, Fasciani A, Casper RF, Varmuza S. Effect of maternal age and conditions of fertilization on programmed cell death during murine preimplantation embryo development. Mol Hum Reprod 1998; 4:139–45 63. Hardy K, Winston RM, Handyside AH. Binucleate blastomeres in preimplantation human embryos in vitro: failure of cytokinesis during early cleavage. J Reprod Fertil 1993; 98:549–58 64. Jamieson ME, Coutts JR, Connor JM. The chromosome constitution of human preimplantation embryos fertilized in vitro. Hum Reprod 1994; 9:709–15 65. Munné S, Alikani M, Tomkin G, Grifo J, Cohen J. Embryo morphology, developmental rates, and maternal age are correlated with chromosome abnormalities. Fertil Steril 1995; 64:382–91 66. Hardy K, Spanos S. Growth factor expression and function in the human and mouse preimplantation embryo. J Endocrinol 2002; 172:221–36

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Figure 9.1 Diagram outlining the main morphological and biochemical features of apoptosis. Reproduced with permission from Hardy K. Apoptosis in the human embryo. Rev Reprod 1999; 4:125–34

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Figure 9.2 Frames from video timelapse of cells blebbing and exploding on the surface of the inner cell mass of an expanded mouse blastocyst

Figure 9.3 Small cytoplasmic blebs (arrowed) on the surface of the inner cell mass of a human blastocyst

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Figure 9.4 Confocal micrograph of a day-4 10-cell embryo with condensed chromatin (arrowed), vacuoles (v) and a binucleate cell (b). Nuclei are labeled with propidium iodide. Reproduced with permission from Hardy K. Apoptosis in the human embryo Rev Reprod 999; 4:125–34

Figure 9.5 Confocal micrograph of a day-6 early human blastocyst with condensed chromatin (arrowed) and a metaphase. Nuclei are labeled with 4′,6-diamidino-2-phenylindole (DAPI)

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Figure 9.6 Three-dimensional confocal reconstruction of nuclei from a day-6 blastocyst showing nucleus with condensed chromatin (arrow), healthy interphase nucleus (i) and a nucleus in mitosis (m). Nuclei are labeled with DAPI. Reproduced with permission from Hardy K. Apoptosis in the human embryo. Rev Reprod 1999; 4:125–34

Figure 9.7 Whole mount of a differentially labeled human blastocyst (labeled as described in reference 19), with the trophectoderm nuclei in

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orange (labeled with propidium iodide and DAPI) and the inner cell mass nuclei in green (labeled with DAPI alone). Note the fragmenting trophectoderm nucleus (arrowed)

Figure 9.8 Two fragmented inner cell mass nuclei (arrowed) from a mouse blastocyst

Figure 9.9 Confocal micrograph of a fragmented nucleus (arrowed) in a day6 human blastocyst with 110 nuclei. Nuclei are labeled with DAPI (highpower view of Figure 9.15)

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Figure 9.10 Confocal micrograph of a day-6 early human blastocyst with 26 nuclei, seven of which are showing clear morphological evidence of apoptosis. The DNA is fragmented in terminal deoxynucleotidyl transferasemediated dUTP nick-end labeled (TUNEL) nuclei (pink) and intact in DAPI-labeled nuclei (blue). Fragmented nuclei with (arrowed) and without (f) TUNEL labeling are clearly shown, as is a condensed nucleus with TUNEL labeling (c), and two nuclei undergoing mitosis (m)

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Figure 9.11 Confocal micrograph of a fragmented, TUNEL-labeled, nucleus (arrowed, pink) and healthy interphase nuclei (green). Reproduced with permission from Hardy K. Apoptosis in the human embryo. Rev Reprod 1999; 4:125–34, and from Hardy K, et al. From cell death to embryo arrest: mathematical models of human preimplantation embryo development. Proc Natl Acad Sci USA 2001; 98:1655–60 © 2001, National Academy of Sciences, USA

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Figure 9.12 Confocal micrograph of a day-4 uncompacted human embryo with 14 cells (and 16 nuclei). Only the polar body is TUNEL-labeled (pink, arrowed), showing DNA fragmentation. Nuclei with intact DNA are labeled with DAPI (blue)

Figure 9.13 Confocal micrograph of a day-5 human morula with 29 nuclei. Two condensed nuclei are TUNELlabeled (pink, arrowed), one fragmented nucleus is TUNEL-labeled (f) and one nucleus is in mitosis (m).

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Nuclei with intact DNA are labeled with DAPI (blue)

Figure 9.14 Confocal micrograph of a day-5 human blastocyst with 82 nuclei. Only one nucleus is TUNEL-labeled (pink, arrowed). Nuclei with intact DNA are labeled with DAPI (blue)

Figure 9.15 Confocal micrograph of a day-6 hatching human blastocyst with 95 nuclei, 11 of which are TUNELlabeled (pink, arrowed). Nuclei with intact DNA are labeled with DAPI (blue)

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Figure 9.16 Confocal micrograph of a day-6 human blastocyst with 89 nuclei, 14 of which are TUNEL-labeled (pink, arrowed). Nuclei with intact DNA are labeled with DAPI (blue)

Figure 9.17 Confocal micrograph of a day-6 human blastocyst with 45 nuclei, and extensive TUNEL-labeling (pink). Nuclei with intact DNA are labeled with DAPI (blue)

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Figure 9.18 Immunohistochemical localization of bcl-2 (brown staining) in a day-5 human blastocyst of poor morphology. Reproduced with permission from Spanos S, et al. Caspase activity and expression of cell death genes during development of human preimplantation embryos. Reproduction 2002; 124:353–63

Figure 9.19 Immunohistochemical localization of bax (brown staining) in a day-5 human blastocyst of poor morphology (same blastocyst as Figure 9.18). Note low level of staining in

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some cells (arrow). Reproduced with permission from Spanos S, et al. Caspase activity and expression of cell death genes during development of human preimplantation embryos. Reproduction 2002; 124:353–63

Figure 9.20 Confocal micrograph of a day-3 compacting human embryo with 15 nuclei showing one cell with active caspases present in the cytoplasm (arrowed, green). Nuclei are labeled with DAPI (red). Reproduced with permission from Spanos S, et al. Caspase activity and expression of cell death genes during development of human preimplantation embryos. Reproduction 2002; 124:353–63

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Figure 9.21 Confocal micrograph of a day-6 expanded human blastocyst with 31 nuclei. Active caspases are green. Note association of active caspases with a fragmented nucleus (arrowed). Nuclei are labeled with DAPI (red). Reproduced with permission from Spanos S, et al. Caspase activity and expression of cell death genes during development of human preimplantation embryos. Reproduction 2002; 124:353–63

Figure 9.22 Early human blastocyst (day-6) with vacuolated excluded cell

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(arrowed) lying between the trophectoderm and the zona pellucida

Figure 9.23 Human blastocyst of poor morphology with excluded cells (arrowed) lying between the trophectoderm and the zona pellucida

Figure 9.24 Hatching human blastocyst with excluded cells (arrowed) lying in the blastocoel cavity. Reproduced with permission from The Society for Endocrinology from Hardy K. Growth factor expression and function in the human and mouse preimplantation embryo. J Endocrinol 2002; 172:221–36

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Figure 9.25 Human embryo (day 3) with extensive fragmentation (arrowed)

Figure 9.26 Cleavage-stage human embryo with large vacuoles in the cytoplasm

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Figure 9.27 Confocal micrograph of day-5 human blastocyst showing engulfed cell (arrow). Reproduced with permission from Hardy K. Apoptosis in the human embryo. Rev Reprod 1999; 4:125–34

10 Human implantation Owen Davis and Zev Rosenwaks

Introduction The process of implantation entails complex interactions between blastocyst and uterine cells (Figure 10.1). Successful implantation can only be initiated when preembryonic development is synchronized with the receptive state of the endometrium. Preparation of the endometrial bed is dependent not only on adequate hormonal stimulation, but also on a reciprocal dialog between the blastocyst and endometrium that is mediated by several factors including cytokines, growth factors and adhesion molecules, which are elaborated by the endometrium and the blastocyst. This chapter provides an overview of currently postulated mechanisms of implantation in the human.

Early embryogenesis (Figure 10.2) Fertilization of the mature, metaphase II oocyte occurs in the ampullary region of the Fallopian tube. A combination of segmental tubal contractions and endosalpingeal ciliary activity transports the preembryo into the uterine lumen at the morula stage, within 72 h of ovulation. Further mitotic cleavage ensues over the next few days while the conceptus remains free within the uterine cavity. Requisite to the process of implantation is the differentiation of the preembryo into a blastocyst, comprising specialized extraembryonic epithelial cells (trophectoderm) and the inner cell mass. All mammalian preembryos must hatch from their zonae pellucidae prior to their initial loose attachment to the endometrial surface (apposition). The process of thinning and degradation of the zona pellucida appears to be primarily driven by the blastocyst, and permits the initial cell membrane contact of the blastocyst polar trophectoderm with the endometrial luminal epithelium. Studies in non-human primates have shown that mononuclear cytotrophoblasts have fused into syncytia prior to attachment of these cells to the endometrial surface1. In the human it is the syncytial trophoblast that first adheres to the endometrial surface and subsequently invades the epithelium during the initial days of implantation (Figure 10.3). Once the blastocyst is completely embedded, cytotrophoblast columns further invade the uterine wall.

The window of implantation Although the human blastocyst’s requirements for nidation may not be rigorous, as evidenced by the occurrence of ectopic gestation, the uterus is receptive to implantation

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only during a discrete interval. The search for molecular markers and/or mediators of the receptive state must therefore focus on this temporal window. In vitro fertilization (IVF) has provided investigators with a useful scientific model for studying the window of transfer (and by extrapolation, implantation) in the human. Here, preembryos of known ‘age’ are transferred into temporally defined endometria. With conventional IVF, however, oocytes are harvested following controlled ovarian hyperstimulation, which typically results in peak luteal phase peripheral estradiol and progesterone levels and estradiol/progesterone ratios far exceeding those seen in the natural cycle. Endometria from pharmacologically stimulated cycles typically display histological and morphological advancement2. A more physiological model for study of the window of transfer is provided by oocyte donation. Preembryos of defined age, derived from in vitro fertilized donated oocytes, are transferred to recipients lacking endogenous ovarian function whose endometria are exposed to physiological levels of estrogen and progesterone. Steroid replacement regimens are specifically designed to mimic the normal pattern of hormone secretion observed in the natural menstrual cycle. Late secretoryphase endometrial biopsies obtained in preparatory hormone-replacement cycles typically demonstrate in-phase histology, while biopsies performed on cycle day 20–21 reveal appropriately developed stroma with lagging glandular histology (day 17–18)3. An analysis of the donor oocyte literature suggests that the window for transfer of 4–8-cell stage preembryos spans endometrial cycle days 16–20. Given the expected time for early cleavage-stage preembryos to develop into blastocysts, this suggests that the window of implantation in the human is approximately 4 days, commencing on cycle day 19–20 and extending to day 24.

Endometrial ultrastructure in the window of implantation Scanning electron microscopy has revealed a striking morphological transformation of the endometrial surface in the perinidatory window. Surface microvilli and cilia are seen to regress and, under the influence of progesterone, bulbous luminal apical protrusions termed ‘pinopods’ develop4 (Figure 10.4). Pinopods are seen in 78% of endometrial biopsies on postovulatory day 6, yet are scarce on postovulatory days 2 and 9. This temporal expression of pinopods suggests that they may play an as yet undetermined functional role in implantation, such as pinocytosis/endocytosis of uterine fluid and/or increasing the surface of blastocyst-epithelial contact. Histological evaluation of secretory endometrium in the window of implantation reveals pronounced stromal edema5. Theoretically, this could serve to facilitate engulfment of the blastocyst. Although these various morphological changes in the endometrium may serve as markers of uterine receptivity, their functional significance remains unclear.

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Cell adhesion molecules Cell adhesion molecules are subdivided into four major subclasses: integrins, cadherins, selectins and members of the immunoglobulin superfamily. Cell adhesion molecules have been implicated in both the cyclic regeneration of the functional layer of the endometrium and blastocyst implantation. Integrins Integrins are ubiquitous heterodimeric transmembrane glycoproteins, comprising noncovalently bound α- and β-subunits. Integrins play a role in cell-cell binding and in cellular interactions with the extracellular matrix. The endometrium expresses both constitutive and cycle-dependent integrins. It should be noted that both endometrial epithelial estrogen and progestin receptors are down-regulated in the mid-luteal phase, during the window of implantation6. The decline of progesterone receptors is temporally associated with the appearance of integrin avb3 on the surface epithelium, which is expressed after day 197. Additionally, αvβ3 has been found covering the preimplantation embryo8 It is theorized that αvβ3 may offer adhesion sites for the blastocyst. Additionally, it has been suggested that αvβ3 binds and activates matrix metalloproteinases and plasminogen activators in the extracellular matrix9, indicating that this integrin may function as both an endometrial receptor for the blastocyst and a facilitator of trophoblastic invasion. Clinical investigation has suggested that mid-luteal avp3 is a marker of human uterine receptivity. A number of disorders associated with infertility have been associated with reduced expression of αvβ3, including endometriosis10, luteal phase defect11, idiopathic infertility12 and the presence of hydrosalpinges13 (which in turn have been associated with impaired implantation following IVF and embryo transfer). Although a functional role for specific integrins has not been established, further research encompassing their clinical utility as markers, or a physiological role as mediators of implantation, is warranted. Mucins As a class, mucins are high-molecular-weight and highly glycosylated molecules found in many secretory epithelia, and specifically on the apical aspect of endometrial epithelial cells. Various mucins may play a role in blastocyst attachment14. In the human, MUC-1 is up-regulated during the implantation window in endometrial samples15. It has been suggested that MUC-1 may regulate blastocyst adhesion, and that the blastocyst, in turn, locally modulates MUC-1 at the implantation site, focally rendering the uterine epithelium adhesive while the remainder remains non-adhesive16. Another mucin which has received attention as a possible marker of human endometrial receptivity is MAG (mouse ascites Golgi). MAG appears on the luminal epithelial surface of the endometrium on cycle day 18–19; abnormal expression of MAG has been cited in some patients with idiopathic infertility17.

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Attachment: the role of growth factors and cytokines Growth factors and cytokines are ubiquitous families of peptides and proteins which variously exhibit paracrine, autocrine and endocrine activity. In the endometrium, specific growth factors, cytokines and their respective receptors and binding proteins demonstrate cycle dependence, and appear to play a role in the dialog between the preembryo and uterus. The epidermal growth factor (EGF) family includes the polypeptides EGF, heparinbinding EGF-like growth factor (HB-EGF), transforming growth factor-α (TGF-α), amphiregulin and betacellulin; all interact with the EGF receptor (EGF-R). Studies in the murine model have demonstrated uterine epithelial expression of HB-EGF around the pre-attachment blastocyst; furthermore, HB-EGF appears to stimulate blastocyst growth and hatching in vitro, via EGF receptors on the blastocyst surface18. Colony-stimulating factor-1 (CSF-1) is a glycosylated homodimer, also referred to as macrophage CSF-1. CSF-1 is preferentially expressed in human endometrial glands during the mid-secretory phase and in first-trimester decidua19. Furthermore, transcripts encoding the CSF-1 receptor have been identified in the preimplantation human embryo20, suggesting a possible role for endometrial CSF-1 in implantation. Leukemia inhibitory factor (LIF) is a glycoprotein displaying both proliferative and differentiative effects through interaction with its receptor. The LIF receptor is composed of two subunits, the LIF receptor (LIF-R) and glycoprotein 130 (gp 130). Blastocyst implantation fails to occur in knock-out mice lacking a functional LIF gene21. In the human, LIF is maximally expressed in mid- and late secretory-phase glandular and luminal epithelium22. LIF appears to regulate human cytotrophoblasts, modulating their differentiation to the anchoring phenotype23. In vitro studies have additionally indicated that LIF regulates human trophoblast differentiation along the invasive pathway, suggesting a role in the invasive phase of implantation24. Clinical studies have indicated that abnormal expression of endometrial LIF may be associated with human infertility. Mutations in the coding region of the LIF gene have been identified in some infertile women25, and a greater proportion of secretory phase uterine flushings have undetectable levels of LIF in women with idiopathic infertility when compared with fertile controls26. The interleukin-1 (IL-1) family comprises two homologous polypeptides (IL-lα and IL-1β), their receptors (IL-1R) and the IL-1 receptor antagonist (IL-1ra). The entire IL-1 system is expressed in human endometrium; IL-1 receptors have been found to increase significantly in the mid-luteal phase27. Human preimplantation embryos also express IL1, IL-1R and IL-lra28. Of note, the presence of IL-1 in preembryo culture fluid is a positive predictor of implantation potential29. Interaction of the blastocyst with receptive endometrium elicits embryonic secretion of IL-1, which in turn effects localized changes in the endometrium prior to adhesion; the embryonic IL-1 system appears to play a role specifically in the up-regulation of endometrial β3 integrin30.

Trophoblast invasion (Figure 10.5) Following adhesion, the trophoblast intrudes, penetrating through the luminal epithelium, reaching and then extending through the basal lamina with subsequent invasion into the

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endometrial stroma, and ultimately invades the maternal vessels with establishment of the hemochorial placenta. The presence of the blastocyst initiates the decidual reaction, wherein stromal cells undergo proliferation and differentiation into specialized cells which may provide nutritional support to the preembryo and play a regulatory role in trophoblast invasion into the stroma. Of note, the decidual reaction can be experimentally induced via stimulation/trauma to the endometrium. The attachment of the invading trophoblast to the extracellular matrix (ECM) is mediated by cell adhesion molecules. It is likely that switching of specific integrins on the trophoblast membranes is one manifestation of the transition of the cytotrophoblasts to the invasive phenotype31. Modulated and localized proteolysis of ECM is another key component of the invasive phase of implantation. First-trimester invading cytotrophoblasts express specific matrix-degrading enzymes such as the matrix metalloproteinases MMP-2 and MMP-932; the latter degrades laminin/type IV collagenrich basement membranes. Of note, IL-1β may play an autocrine role in the promotion of cytotrophoblast invasiveness via stimulation of MMP-9 production33. Other proteinases, such as urokinase-type plasminogen activator, may also facilitate degradation of the ECM by the invading trophoblast, either directly or via activation of pro-MMPs34. The activity of the various proteinases is in turn modulated by specific protease inhibitors of both decidual and trophoblastic origin. These inhibitors appear to play an important role in limiting invasion. A number of tissue inhibitors of metalloproteinases (TIMPs) have been identified. TIMP-3 is expressed in both endometrial stroma and trophoblasts, and regulates the activity of MMP-935. IL-1b has been observed to inhibit the activity of TIMP-3 in human decidua, indicating that the trophoblast can promote its own invasiveness through the inhibition of maternal restraint mechanisms36. Specific ECM proteins also play a role in implantation. Decidualized endometrial stromal cells elaborate both laminin and fibronectin. Laminin may play a permissive role in invasion; in vitro studies have shown that human trophoblasts readily attach to laminin-coated surfaces37. Conversely, fibronectin appears to promote maternal restraint through the inhibition of cytotrophoblast invasion. Other factors have additionally been implicated in the maternal inhibition of trophoblastic invasion. Insulin growth factor binding protein-1 (IGFBP-1) is abundantly produced in secretory endometrium and decidua. IGFBP-1 has been demonstrated to bind human cytotrophoblast via α5β1 integrin in the cell membrane, and restrains cytotrophoblast invasion in endometrial stromal culture38. Transforming growth factor-β1 (TGF-β1) also appears to be an effector of maternal restraint on trophoblast invasion. TGF-β1 is abundantly expressed in maternal endometrium, and exerts several modulatory effects, including: inhibition of cytotrophoblast proliferation and promotion of differentiation into syncytiotrophoblasts, and induction of cytotrophoblast protease inhibitors such as TIMP-1 and plasminogen activator inhibitor (PAI)39.

Conclusion The successful propagation of the human species depends on reproductive efficiency. Implantation, a critical component of reproductive physiology, remains one of the most incompletely understood processes in reproductive biology. Implantation involves the

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introduction of a competent blastocyst into the uterine lumen during a temporally restricted interval when ovarian hormones induce a receptive uterine phase. Reciprocal signalling between the blastocyst and the endometrium is mediated in some measure by cytokines, growth factors and adhesion molecules. Successful ‘cross-talk’ leads to apposition and attachment of the blastocyst to the uterine wall, followed by intrusion and invasion. The process of trophoblastic invasion entails a delicate interplay between invasion facilitators and invasion inhibitors, i.e. promotion and restraint. Further elucidation of the complex mechanisms underlying the process of implantation in the human should provide clinical inroads into the diagnosis and treatment of reproductive failure.

References 1. Enders AC. Current topic: structural responses of the primate endometrium to implantation. Placenta 1991; 12:309–25 2. Martel D, Frydman R, Sarantis L, Roihe D, Psychoyos A. Scanning electron microscopy of the uterine luminal epithelium as a marker of the implantation window. In Yoshinaga K, ed. Blastocyst Implantation. Boston: Serono Symposia, 1989:225–30 3. Rosenwaks Z. Donor eggs: their application in modern reproductive technologies. Fertil Steril 1987; 47:895–909 4. Martel D, Monier MN, Roiche D, Psychoyos A. Hormonal dependence of pinopode formation at the uterine luminal surface. Hum Reprod 1991; 6:597–603 5. Noyes RW, Hertig AT, Rock J. Dating the endometrial biopsy. Fertil Steril 1950; 1:3–25 6. Lessey BA, Killam AP, Metzger DA, et al. Immunohistochemical analysis of human uterine estrogen and progesterone receptors throughout the menstrual cycle. J Clin Endocrinol Metab 1988; 647:334–40 7. Lessey BA, Damjanovich L, Coutifaris C, et al. Integrin adhesion molecules in human endometrium: correlation with normal and abnormal menstrual cycles. J Clin Invest 1992; 90:188–95 8. Campbell S, Swann HR, Seif MW, et al. Cell adhesion molecules on the oocyte and preimplantation human embryo. Hum Reprod 1995; 10:1571–8 9. Brooks PC, Stromblad S, Sanders LC, et al. Localization of matrix metalloproteinase MMP-2 to the surface of invasive cells by interaction with integrin αvβ3. Cell 1996; 85:683–93 10. Lessey BA, Castelbaum AJ, Sawin SW, et al. Aberrant integrin expression in the endometrium of women with endometriosis. J Clin Endocrinol Metab 1994; 79:643–9 11. Lessey BA, Yeh I, Castelbaum AJ, et al. Endometrial progesterone receptors and markers of uterine receptivity in the window of implantation. Fertil Steril 1996; 65:477–83 12. Lessey BA, Castelbaum AJ, Sawin SW, et al. Integrins as markers of uterine receptivity in women with primary unexplained infertility. Fertil Steril 1995; 63:535–42 13. Meyer WR, Castelbaum AJ, Somkuti S, et al. Hydrosalpinges adversely affect markers of endometrial receptivity. Hum Reprod 1997; 12:1393–8 14. Carson DD, Rohde LH, Surveyor G. Cell surface glycoconjugates as modulators of embryo attachment to uterine epithelial cells. Int J Biochem 1994; 26:1269–77 15. Hey NA, Graham RA, Seif MW, et al. The polymorphic epithelial mucin MUC 1 in human endometrium is regulated with maximal expression in the implantation phase. J Clin Endocrinol Metab 1994; 78:337–42 16. Meseguer M, Aplin JD, Caballero-Campo P, et al. Human endometrial mucin MUC1 is upregulated by progesterone and down-regulated in vitro by the human blastocyst. Biol Reprod 2001; 64:590–601

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17. Kliman HJ, Feinberg RJ, Schwartz LB, et al. A mucin-iike glycoprotein identified by MAG (mouse ascites Golgi) antibodies. Menstrual cycle-dependent localization in human endometrium. Am J Pathol 1995; 146:166–81 18. Das SK, Wang X, Paria BC, et al. Heparin-binding EGF-like growth factor gene is induced in the mouse uterus temporally by the blastocyst solely at the site of its apposition: a possible ligand for interaction with blastocyst EGF-receptor in implantation. Development 1994; 120:1071–83 19. Bartocci A, Pollard JW, Stanley ER. Regulation of CSF-1 during pregnancy. J Exp Med 1986; 164:956–61 20. Sharkey AM, Dellow K, Blayney M, et al. Stage-specific expression of cytokine and receptor messenger ribonucleic acids in human preimplantation embryos. Biol Reprod 1995; 53:974–81 21. Stewart CL, Kaspar P, Brunet LJ, et al. Blastocyst implantation depends on maternal expression of leukaemia inhibitory factor. Nature (London) 1992; 359:76–9 22. Charnock-Jones DS, Sharkey AM, Fenwick P, et al. Leukaemia inhibitory factor mRNA concentration peaks in human endometrium at the time of implantation and the blastocyst contains mRNA for the receptor at this time. J Reprod Fertil 1994; 101:421–6 23. Nachtigall MJ, Kliman HJ, Feinberg RF, et al. The effect of leukemia inhibitory factor (LIF) on trophoblast differentiation: a potential role in human implantation. J Clin Endocrinol Metab 1996; 81:801–6 24. Bischof P, Haenggeli L, Campana A. Effect of leukemia inhibitory factor on human cytotrophoblast differentiation along the invasive pathway. Am J Reprod Immunol 1995; 34:225–30 25. Giess R, Tanasescu I, Steck T, et al. Leukaemia inhibitory factor gene mutations in infertile women. Mol Hum Reprod 1999; 5:581–6 26. Laird SM, Tuckerman EM, Dalton CF, et al. The production of leukaemia inhibitory factor by human endometrium: presence in uterine flushings and production by cells in culture. Hum Reprod 1997; 12:569–74 27. Simon C, Piquette GN, Frances A, et al. Localization of interleukin-1 type-1 receptor and interleukin-1β in human endometrium throughout the menstrual cycle. J Clin Endocrinol Metab 1993; 77:549–55 28. Krussel JS, Simon C, Rubio MC, et al. Expression of interleukin-1 system mRNA in single blastomeres from human preimplantation embryos. Hum Reprod 1998; 13:2206–11 29. Sheth KV, Roca GL, al-Sedairy ST, et al. Prediction of successful embryo implantation by measuring interleukin 1α and immunosuppressive factor(s) in preimplantation embryo culture fluid. Fertil Steril 1991; 55:952–7 30. Simon C, Gimeno MJ, Mercader A, et al. Embryonic regulation of integrins β3, α4 and α1 in human endometrial epithelial cells in vitro. J Clin Endocrinol Metab 1997; 82:2607–16 31. Damsky CH, Librach C, Lim KH, et al. Integrin switching regulates normal trophoblast invasion. Development 1994; 120:3657–66 32. Bischof P, Friedli E, Martelli M, et al. Expression of xtracellular matrix-degrading metalloptroteinases by cultured human cytotrophoblast cells: effects of cell adhesion and immuno-purification. Am J Obstet Gynecol 1991; 165:1791–801 33. Librach CL, Feigenbaum SL, Bass KE, et al. lnterleukin-1(3 regulates human cytrotrophoblast metalloproteinase activity and invasiveness in vitro. J Biol Chem 1994; 269:17125–31 34. Queenan JT Jr, Kao LC, Arboleda A, et al. Regulation of urokinase-type plasminogen activator production by cultured human cytotrophoblasts. J Biol Chem 1987; 262:10903–6 35. Higuchi T, Kanzaki H, Nakayama H, et al. Induction of tissue inhibitor of metalloproteinase 3 gene during in vitro decidualization of human endometrial stromal cells. Endocrinology 1995; 136:4973–81 36. Huang HY, Wen Y, Irwin JC, et al. Cytokine mediated regulation of tissue inhibitor of metalloproteinase-1 (TIMP-1), TIMP-3, and 92-kDa type IV collagenase mRNA expression in human endometrial stromal cells. J Clin Endocrinol Metab 1998; 83:1721–9

An atlas of human blastocysts 284 37. Loke YW, Gardner L, Burland K, et al. Laminin in human trophoblast-decidua interaction. Hum Reprod 1989; 4:457–63 38. Irwin JC, Giudice LC. IGFBP-1 binds to the a5b1 integrin in human cytotrophoblasts and inhibits their invasion into decidualized endometrial stromal cells in vitro. Growth Horm IGF Res 1998; 8:21–31 39. Graham CH, McCrae KR, Lala PK. Molecular mechanisms of controlling trophoblast invasion of the uterus. Trophoblast Res 1993; 7:237–50

Figure 10.1 Implantation involves complex interactions between blastocyst and uterine cells. Reprinted with permission from Moore KL, Persaud TVN, Shiota K, eds. Color Atlas of Clinical Embryology. Philadelphia: WB Saunders, 1994:5

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Figure 10.2 Timetable of early embryogenesis. Schematic courtesy of Keith L.Moore, The Developing Human: Clinically Oriented Embryology, 3rd edn. Philadelphia: WB Saunders, 1982

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Figure 10.3 Trophoblastic projections. As observed in this sequence of timelapse photographs, the hatched human blastocyst produces small projections that exhibit amoeboid movement, appear fimbriated and move in a fashion that suggests they seek an

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implantation site. Sequence courtesy of L.Veeck

Figure 10.4 Pinopods. Luminal epithelium of the human uterus on day 6 after ovulation during spontaneous menstrual cycle. Courtesy of D.Martel2. In Koji Yoshinaga, ed. Blastocyst Implantation. Boston: Serono Symposia, 1989:228

Figure 10.5 Implanting human blastocyst. The blastocyst securely anchors itself in the endometrium; to the right, veritable cables are visible. Courtesy of Lennart Nilsson/Albert Bonniers Forlag AB, A Child is Born, Dell Publishing Company

11 Human embryonic stem cells Michal Amit and Joseph Itskovitz-Eldor

Introduction Embryonic stem (ES) cells are continuously growing cell lines of embryonic origin, first isolated from the inner cell mass (ICM) of mouse blastocysts1,2. These unique cells are characterized by the following features: (1) They are derived from preimplantation embryos; (2) In culture, they indefinitely maintain uniform colonies of undifferentiated morphology, i.e. high nucleus/cytoplasm ratio and the presence of two to four nucleoli; (3) They form teratomas after injection into SCID (severe combined immunodeficiency) mice, proving their potential to form all three embryonic germ layers; (4) They maintain normal karyotype after continuous culture. These features are summarized in Table 11.1. Given their ability to differentiate into any cell type of the adult body, it is not surprising that much effort was invested in the development of human ES cell lines. Since the first publication on the derivation of human pluripotent cell lines3, additional lines have been reported that meet the ES cell criteria4–6. According to a list published by the National Institutes of Health (NIH, www.nih.gov/news/stemcell/index.htm), there are more than 70 human ES cell lines in several laboratories around the world that fulfil the listed features (Table 11.1) of ES cells. The availability of dozens of cell lines suggests that the derivation of these lines is a reproducible procedure with reasonable success rates.

Derivation of human ES cell lines Human ES cell lines may be derived using two alternative methods, namely immunosurgical or mechanical isolation of the ICM. In both cases, surplus blastocysts donated by couples undergoing in vitro

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Table 11.1 Main features of embryonic stem (ES) cells (1) Derived from preimplantation embryos (2) Pluripotent, capable of differentiating into representative cells from all three germ layers of the embryo (3) Immortal, with long-term proliferative ability at the undifferentiated stage (self-maintenance) and high telomerase activity (4) Express unique markers typical of cells at the undifferentiated stage, such as transcription factor Oct-4, or cell surface markers such as stage-specific embryonic antigen-3 (SSEA3), SSEA4 and tumor-rejecting antigen-60 (TRA-1–60), TRA1-81 (5) Maintain normal karyotype after prolonged culture (6) Clonogenic, i.e. each individual cell possesses these characteristics

fertilization (IVF) treatments are grown in vitro to the blastocyst stage (Figure 11.1). The first method was developed in the early 1970s by Solter and Knowles, to derive embryonic teratocarcinoma (EC) lines and for early embryonic development research7. During immunosurgery, the outer layer of the trophectoderm is selectively removed, leaving an intact ICM. Initially, the zona pellucida of the embryo is removed using ether Tyrode’s solution or pronase, after which the embryo is exposed to anti-human whole antiserum (Figure 11.2a). These antibodies recognize and attach to any human cell, thereby marking all trophoblast cells. As cell-cell connections between the trophoblast cells prevent anti-body penetration into the embryo, the ICM of the embryo remains intact. The next stage includes exposure of the embryo to guinea-pig complement, which lyses all cells marked with the antibody, i.e. the trophectoderm cells (Figure 11.2b). The intact ICM is further cultured on a mitotically inactivated mouse embryonic fibroblast (MEF) feeder layer (Figure 11.2c). Human ES cell lines may also be derived using mechanical separation of the trophectoderm and from ICM. The latter may be conducted by either removing the trophectoderm using 27-gauge needles followed by plating the ICM on MEFs, or plating an intact blastocyst on MEFs, culturing until the ICM cells expand and then mechanically removing the ICM cells into a fresh culture plate. MEF feeder layers perform a dual role: they support ES cell growth and prevent their spontaneous differentiation during culture. The mechanism by which MEFs prevent differentiation is still not completely understood. Six human ES cell lines (I-3, I-4, I-6, I-8, J-6 and J-3) have been derived in our laboratory that fulfil the characteristics of existing human pluripotent cell lines. These lines demonstrate typical morphology of human ES cell colonies (illustrated in Figure 11.3) and single cells, i.e. high nucleus/cytoplasm ratio and the presence of at least two nucleoli (shown in Figure 11.4). In addition, scanning electron microscopy reveals the existence of cytoplasmic connections between human ES cells and MEF(Figure 11.5). Compared with mouse ES cells, primate ES cells have been found to express different surface markers, specific to undifferentiated cells. While mouse ES cells highly express

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surface marker stage-specific embryonic antigen-1 (SSEA1), and do not express SSEA3, SSEA4, tumor-rejecting antigen-60 (TRA-1–60) and TRA-1–81, non-human primate ES cells and human ES cells strongly express SSEA4, TRA-1–60 and TRA-1–81, weakly express SSEA3 and do not express SSEAl3,4. The typical antibody expression of human ES cells is demonstrated in Figure 11.6.

Human ES cell culture At first, human ES cells required an MEF feeder layer to grow continuously at the undifferentiated stage. Mouse ES cells can be grown as undifferentiated cells directly on gelatin-coated plates with the addition of leukemia inhibitory factor (LIF). Unfortunately, LIF does not have the same effect on human ES cells3,4. Three major improvements in the basic culture methods of human ES cells have been developed since their original derivation on an MEF feeder layer: (1) The use of serum-free medium, which provides better defined culture conditions8; (2) The ability to grow and derive human ES cells with human feeder layer9; (3) The ability of human ES cells to grow under feeder-free culture conditions10. Human ES cells can be grown as undifferentiated cells using serum replacement supplemented with 4 ng/ml basic fibroblast growth factor (bFGF) while maintaining all ES cell features8. Under these culture conditions, the morphology of human ES cell colonies is slightly different: the cells are organized in more than one layer, while in fetal bovine serum (FBS) the colonies consist of a monolayer of ES cells. Representative examples of colonies in these two conditions are illustrated in Figure 11.7. The serumfree growth of human ES cells leads to better-defined culture conditions for the growth and manipulation of these cells. Another advantage of the serum-free condition is that it has been found to be suitable for the derivation of single-cell human ES cell clones8. As mentioned above, human ES cell lines are derived from the ICM. The ICM cells may not represent a homologous cell population; therefore, it is possible that the pluripotency of human ES cells reflects the combined developmental potential of this cell population. However, the ability of singlecell clones to differentiate into representative tissues of the three embryonic germ layers eliminates this possibility. To date, nine single-cell clones from six various parental ES cell lines have been derived in our laboratory using serum-free conditions. A clonally derived human ES cell line in a 96-well plate is shown in Figure 11.8. Human ES cells have also been reported to grow in an animal-free system9. The suggested animal-free culture system for human ES cells consists of coculture with human fetal-derived feeder layers or human adult Fallopian tube epithelial feeder layers, using medium supplemented with human serum. These conditions are suitable both for the maintenance of their characteristics during prolonged culture and for the derivation of new human ES cell lines. We have demonstrated that foreskin feeder layers support the growth of human ES cells in the above serum-free conditions11. After more than 70 passages, human ES cell lines grown on foreskin feeders exhibited all human ES cell features, including teratoma and embryoid body (EB) formation, expression of surface markers typical of undifferentiated cells and preservation of normal karyotypes. The

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morphology of a human ES cell colony grown on a foreskin line varies from that of a colony grown on MEF in the tendency of the cells to organize in an elliptical manner (demonstrated in Figure 11.9). Unlike fetal fibroblasts, which can grow to reach a certain limited passage, foreskin fibroblasts can grow to reach passage 42. Therefore, foreskin lines have an advantage when large-scale growth of human ES cells is concerned. Lastly, a suggested culture system in which human ES cells are grown on matrigel, laminin or fibronectin, using 100% MEF-conditioned medium supplemented with serum replacement and bFGF, has been reported10. Examples of human ES cells grown using this system for 12 passages are illustrated in Figure 11.10. Since human ES cells may still be exposed to animal pathogens through the conditioned medium, and there is still a requirement for simultaneous massive growth of MEFs for the production of conditioned medium, the quest for the ideal animal- and feeder layer-free culture system for human ES cells is still taking place.

Human ES cell differentiation As long as human ES cells are grown on an MEF feeder layer and passaged routinely every 4–6 days, they will maintain their pluripotency and remain at the undifferentiated stage. They differentiate spontaneously when grown to confluency on MEFs or removed from the MEF feeder layer, and, when plated as crowded cultures on gelatin, they differentiate rapidly and form visible structures (Figure 11.11). Another way to encourage spontaneous differentiation of human ES cells is by inducing the formation of EBs. Human ES cells, like mouse ES cells, spontaneously create EBs when cultured in suspension12. Initially, after 24 h in suspension, they spontaneously form small cell aggregates (Figure 11.12a). These aggregates tend to organize into a special structure consisting of an internal ectoderm layer and an external endoderm layer (Figure 11.12b). Later on, some of the EBs form cysts (Figure 11.12c and d) and visible structures (Figure 11.12e and f). These EBs contain derivatives of the three embryonic germ layers12. Examples of different cell types resulted in differentiating ES cells in EBs are shown in Figure 11.13. It seems that the formation of EBs encourages ES cells to differentiate and consequently increase the rate and efficiency of differentiation. Human ES cells can create EBs with the same efficiency as mouse ES cells, although EBs formed from human ES cells have been found to be somewhat less organized than those derived from mouse ES cells12. The simplest and most convincing way to examine ES cell pluripotency is to look for teratoma formation. Following injection into the hind leg muscle of SCID mice, ES cells spontaneously create teratomas, in which they differentiate into representative tissues of the three embryonic germ layers. Several examples of resultant teratomas formed by lines 1–3 and 1–6 are illustrated in Figure 11.14: from endodermal-origin columnar epithelium, and tubules interspersed with structures resembling fetal glomeruli, from mesodermal-origin mesenchymal tissue, and hyaline cartilage, and from ectodermalorigin epithelium containing melanin-producing cells and nerve tissues. While in EBs, ES cells differentiate mainly into simple structures or unorganized groups of cells. In teratomas, however, human ES cells can also create more complex and

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well-organized organ-like structures. Examples of such organs are illustrated in Figure 11.15, which shows a group of hair follicles, a salivary gland and skin tissue. In their 4 years of existence, human ES cells have been shown to differentiate into specific cell types representative of the three embryonic germ layers in both spontaneous and directed-differentiation models. These differentiation models are summarized in Table 11.2. Overall, the reports on human ES cell differentiation models support the possibility of creating a well-developed model of differentiation for human ES cells, which would resemble the existing mouse ES cell differentiation models.

Table 11.2 Cell types formed by spontaneous and induced human ES cell differentiation Cell types

Reference

Spontaneous differentiation Cardiomyocytes

Kehat et al.13 (2001),

Endothelial cells

Levenberg et al.14 (2002)

Insulin-secreting cells

Assady et al.15 (2001)

Directed differentiation Neuronal cells

Carpenter et al.16 (2001) Reubinoff et al.17 (2001) Zhang et al.18 (2001) Schuldiner et al.19 (2002)

Trophoblast cells

Xu et al.20 (2002)

Cardiomyocytes

Xu et al.21 (2002)

Acknowledgements The authors wish to thank Mrs Hadas O’Neill for editing the manuscript, Mrs Ruth Tal for editing the figures, Mrs Kohava Shariki and Mrs Victoria Marguletz for technical assistance, and Dr D.Levanon and Mrs Zelda Weinberg for their assistance with the electron microscopy work. The antibodies used for the surface marker analysis (Figure 11.6) were generously provided by Professor P.W.Andrews from the University of Sheffield. The research performed in our laboratory was partly supported by the Fund for Medical Research and Development, Rambam Medical Center, and Technion Research and Development Fund, Haifa, Israel.

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References 1. Evans MJ, Kaufman MH. Establishment in culture of pluripotential cells from mouse embryos. Nature (London) 1981; 292:154–6 2. Martin GR. Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci USA 1981; 78:7634–8 3. Thomson JA, Itskovitz-Eldor J, Shapiro SS, et al. Embryonic stem cell lines derived from human blastocysts. Science 1998; 282:1145–7 (erratum in Science 1998; 282:1827) 4. Reubinoff BE, Pera MF, Fong C, Trounson A, Bongso A. Embryonic stem cell lines from human blastocysts: somatic differentiation in vitro. Nat Biotechnol 2000; 18:399–404 5. Lanzendorf SE, Boyd CA, Wright DL, Muasher S, Oehninger S, Hodgen GD. Use of human gametes obtained from anonymous donors for the production of human embryonic stem cell lines. Fertil Steril 2001; 76:132–7 6. Amit M, Itskovitz-Eldor J. Derivation and spontaneous differentiation of human embryonic stem cells. J Anat 2002; 200:225–32 7. Solter D, Knowles BB. Immunosurgery of mouse blastocyst. Proc Natl Acad Sci USA 1975; 72:5099–102 8. Amit M, Carpenter MK, Inokuma MS, et al. Clonally derived human embryonic stem cell lines maintain pluripotency and proliferative potential for prolonged periods of culture. Dev Biol 2000; 227:271–8 9. Richards M, Fong CY, Chan WK, Wong PC, Bongso A. Human feeders support prolonged undifferentiated growth of human inner cell masses and embryonic stem cells. Nat Biotechnol 2002; 20:933–6 10. Xu C, Inokuma MS, Denham J, et al. Feeder-free growth of undifferentiated human embryonic stem cells. Nat Biotechnol 2001; 19:971–4 11. Amit M, Margulets V, Segev H, et al. Human feeder layers for human embryonic stem cells. Biol Reprod 2003; 10:1095 12. Itskovitz-Eldor J, Schuldiner M, Karsenti D, et al. Differentiation of human embryonic stem cells into embryoid bodies comprising the three embryonic germ layers. Mol Med 2000; 6:88– 95 13. Kehat I, Kenyagin-Karsenti D, Snir M, et al. Human embryonic stem cells can differentiate into myocytes with structural and functional properties of cardiomyocytes. J Clin Invest 2001; 108:407–14 14. Levenberg S, Golub JS, Amit M, Itskovitz-Eldor J, Langer R. Endothelial cells derived from human embryonic stem cells. Proc Natl Acad Sci USA 2002; 99:4391–6 15. Assady S, Maor G, Amit M, Itskovitz-Eldor J, Skorecki KL, Tzukerman M. Insulin production by human embryonic stem cells. Diabetes 2001; 50:1691–7 16. Carpenter MK, Inokuma MS, Denham J, Mujtaba T, Chiu CP, Rao MS. Enrichment of neurons and neural precursors from human embryonic stem cells. Exp Neurol 2001; 172:383–97 17. Reubinoff BE, Itsykson P, Turetsky T, et al. Neural progenitors from human embryonic stem cells. Nat Biotechnol 2001; 19:11340–40 18. Zhang S-C, Wernig M, Duncan ID, Brüstle O, Thomson JA. In vitro differentiation of transplantable neural precursors from human embryonic stem cells. Nat Biotechnol 2001; 19:1129–33 19. Schuldiner M, Elges R, Eden A, Yanuka O, Itskovitz-Eldor J, Goldstein RS, Benvisty N. Induced neuronal differentiation of human embryonic stem cells. Brain Res 2001; 913:201–5 20. Xu RH, Chen X, Li DS, et al. BMP4 initiates human embryonic stem cell differentiation to trophoblast. Nat Biotechnol 2002; online 11 November 21. Xu C, Police S, Rao N, Carpenter MK. Characterization and enrichment of cardiomyocytes derived frorn human embryonic stem cells. Circulation Res 2002; 91:501–8

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Figure 11.1 Derivation of human ES cell lines. Preimplantation development of human embryo. In order to produce human ES cell lines, surplus donated human embryos are grown in vitro from early embryos to the blastocyst stage. The ICM is selectively removed and further cultured on mitotically inactivated MEF. The resulting ES cells may

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differentiate into representative tissues of the three embryonic germ layers

Figure 11.2 Derivation of human ES cell lines. Immunosurgery of human

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blastocyst for the derivation of human ES cell lines. (a) Human blastocyst after zona pellucida removal by Tyrode’s solution, during exposure to rabbit anti-human whole antiserum; (b) embryo after exposure to guinea pig complement; (c) intact inner cell mass immediately after immunosurgery on mitotically inactivated mouse embryonic fibroblast feeder layer. Bar 50 µm. From Amit and Itskovitz-Eldor, J Anat 2002; 200:225–32

Figure 11.3 Characterization of human ES cells. Human ES cell colony from line I-4 grown for 47 passages on

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mitotically inactivated MEF. (a) Using low magnification: note the round shape of the colony, typical of human ES cells grown on mitotically inactivated MEF. Bar 100 µm. (b) Using high magnification: typical spaces between the ES cells and high nucleus-to-cytoplasm ratio. Bar 50 µm

Figure 11.4 Characterization of human ES cells. Electron microscopy of single human ES cells from line H-9. Note the high nucleus-to-cytoplasm ratio and the presence of at least two nucleoli. Bar 8 µm

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Figure 11.5 Characterization of human ES cells. Scanning electron microscopy of (a) MEF feeder layer. Note the cytoplasmatic extensions between the cells; (b) H-9.2 cells grown on MEF. Note the cytoplasmatic extension generated by both the ES cells and the MEF. Bar 10 µm

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Figure 11.6 Characterization of human ES cells. Fluorescent immunostaining of human ES cells. (a) Immunostaining of human ES cell colonies I-3 with anti-TRA-1–60 (tumor-rejecting antigen-60) antibodies (×5); (b) immunostaining of human ES cell colony I-6 with anti-SSEA4 (stagespecific embryonic antigen-4) antibodies (×20); (c) immunostaining of human ES cell colony I-6 with antiTRA-1–81 (×20); (d) high-power picture from the marked area in (c) (×63). From Amit and Itskovitz-Eldor, J Anat 2002; 200:225–32

Figure 11.7 Human ES cell culture. Typical morphology of ES cell colonies of clone H-9.2 in different

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types of medium. (a) Medium supplemented with FBS; (b) medium supplemented with serum replacement and bFGF. Bar 50 µm

Figure 11.8 Human ES cell culture. A clonally derived human ES cell line in a 96-well plate. (a) Colony of singlecell clone I-3.3 7 days post-cloning; (b) the same colony at day 12 postcloning. Bar 38 µm

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Figure 11.9 Human ES cell culture. Typical morphology of ES cell colony grown on human foreskin feeder layer. (a) The common morphology: ellipticlike colony of line I-6 after 57 passages on the human feeders, and (b) a rare morphology, irregular shape of line I-3 after 21 passages on the human feeders. Bar 50 µm

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Figure 11.10 Human ES cell culture. Three examples of the colonial morphology of undifferentiated ES cells from clone I-3.2, grown for 12 passages on Matrigel® (BD

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Biosciences, Bedford, MA, USA) matrix using MEF-conditioned medium. Note the typical spaces between the ES cells and the high nucleus-to-cytoplasm ratio ((a) and (c)), and small and dense cells in (b) typical of dividing cells. Bar 50 µm

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Figure 11.11 Human ES cell differentiation in vitro. Structure created by human ES cells H-9 at passage 33, grown to confluency on gelatin (a) for 9 days: note the cyst created by the differentiating ES cells,

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whose structure resembles that of an embryoid body (EB), bar 100 µm; (b) for 10 days: rosettes of neural epithelium-like structures have been formed, bar 75 µm; (c) for 13 days, bar 100 µm

Figure 11.12 Human ES cell differentiation in vitro. Embryoid body (EB) development. (a) EB formed by cells H-9 after 24 h in suspension. Bar 100 µm; (b) EB from line I-6 after 48 h in suspension. Note the ectoderm layer (arrow) surrounding the EB. Bar 75 µm; (c) EB formed by line H-9 starts to create a cyst after 4 days in

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suspension. Bar 50 µm; (d) EB formed by line H-9 after 6 days in suspension. Note the well-developed cyst (white arrows) and some melanin-containing cells (black arrow). Bar 75 µm; (e) and (f) 1-month-old EB from single-cell clones H-9.2 and H-9.2.4, respectively, containing notable structures (white arrows). Bar 50 µm

Figure 11.13 Human ES cells differentiation in vitro. Examples of 1µm epon sections of 1-month-old EB from single-cell clone H-9.2 stained in alkaline toluidine blue. (a) Ball of epithelial-like cells with large regular nuclei and very pronounced nucleoli,

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which indicate intense synthetic activities. Isolated cells are seen with many cytoplasmic lipid droplets. Bar 10 µm; (b) stratified epithelium lining a tube-like structure in which aggregates of unidentified cells can be observed. Bar 25 µm; (c) differentiation of epithelial cells lining a tube-like structure. The surrounding cells are quite varied and probably develop connective tissue cells and intracellular matrices. Bar 25 µm; (d) epithelial-like cells arranged in a solid ball. The cells contain cytoplasmic granular material, which is stained with a deep purple color. Bar 25 µm; (e) stratified epithelium at the periphery of an EB and underlying mesenchymal-like cells of developing connective tissue. Bar 10 µm; (f) development of columnar epithelium seen in a differentiating structure within an EB. Bar 10 µm

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Figure 11.14 Human ES cells differentiation in vivo. Differentiated human ES cells from lines I-3 and I-6 in teratomas. (a) Columnar-epithelium, I-3 (H&E) Bar 25 µm; (b) tubules interspersed with structures resembling fetal glomeruli, I-6 (H&E) Bar 25 µm; (c) mesenchymal tissue, I-6 (H&E) Bar 25 µm; (d) hyalinic cartilage, I-3 (H&E) Bar 100 µm; (e) epithelium containing melanin-producing cells, I6 (H&E) Bar 25 µm; (f) nerve tissue and blood vessels, I-6 (H&E). Bar 50 µm

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Figure 11.15 Differentiated human ES cells from subclone H-9.2.4 in teratomas. (a) Mixed salivary gland with serous and mucous secretory cells

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(H&E); (b) group of developing hair follicles (H&E); (c) complete skin tissue formed by line J-3. Keratincontaining cells facing a large lumen. Bar 50 µm. (a) and (b) taken from reference 6, with permission

12 The mammalian blastocyst as an experimental model Shoukhrat M.Mitalipov, Hung-Chih Kuo and Don P.Wolf

Introduction In the development and application of assisted reproductive technology (ART), clinical experience in the human has often led the way. However, when invasive experimentation is required, the rodent or non-human primate may be the only suitable model. Furthermore, it can be argued that all ART applications should ideally be first applied in animals. Recent advances in non-human primate ART, with emphasis on experiments that cannot be conducted in humans for ethical reasons, are highlighted here. Preimplantation development of the mammalian embryo incorporates explicit developmental stages from the formation of the zygote after fertilization to cleavage, morula formation, compaction and, finally, cavitation with the formation of the blastocyst (Figure 12.1). In vitro, in the rhesus monkey, preimplantation development is slower than in the human, with early blastocysts containing 100 or so cells seen on day 6–7 instead of day 5. Admittedly, in vivo rates are probably somewhat faster than in vitro, which may reflect suboptimal culture conditions. During preimplantation development there is little or no growth in embryo volume, rather the large oocyte simply cleaves into smaller and smaller cells until blastulation. After fertilization, the embryonic genome is transcriptionally inactive and the first divisions of the embryo are supported primarily by maternally inherited proteins and mRNAs present in the cytoplasm of the unfertilized oocyte. The onset of embryonic transcription at the maternal-embryonic transition is species-dependent, starting as early as the 1–2-cell stage in the mouse1 and the 4–8-cell stage in the rhesus monkey and humans2–4. Cavitation (early blastocyst formation) is driven by the expression of specific sets of gene products, including critical gene families: the E-cadherin-catenin cell adhesion family, the tight junction gene family, the Na/K-adenosine triphosphatase (ATPase) gene family and perhaps the aquaporin gene family5. During early preimplantation development, individual blastomeres are totipotent, that is, each blastomere is capable of supporting a pregnancy and the development of a viable fetus. Blastocyst formation marks the first obvious differentiation of the mammalian preimplantation embryo when two distinctive cell types are present: the inner cell mass (ICM) and the trophectoderm. The trophectoderm comprises a sphere of single-layered flat epithelial cells surrounding a fluid-filled cavity, the blastocoel. A compact group of cells within the sphere represents the ICM, whose formation is dependent upon expression of the transcription factor Oct4. During development, trophectoderm and ICM cells contribute to distinctive embryonic lineages. Experimental studies in the mouse

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indicate that the ICM gives rise to the embryo proper and to several extraembryonic membranes, while the trophectoderm contributes to the trophoblast layers of the placenta6. Developmental competence to the blastocyst stage is often used as assessment of embryo viability in the context of infertility treatments such as in vitro fertilization/intracytoplasmic sperm injection (IVF/ICSI), or in the creation of reconstructed embryos by nuclear transfer. To be sure, however, in vitro development to the blastocyst stage does not ensure full-term development, since all blastocysts are not the same. For instance, parthenotes reach the blastocyst stage but do not develop beyond the 25-somite stage7. Even after fertilization, significant differences in blastocyst quality have been observed, and it is recognized that morphological assessment alone is not an absolute measure of viability8. The ultimate measure of embryo quality is unquestionably the ability to support full-term pregnancy following transfer to a recipient. Unfortunately, it is often impractical to use term birth as the outcome measure because of time and resource considerations. On the other hand, the embryo’s ability to grow to the blastocyst stage is a significant accomplishment, as progression beyond the maternal-embryonic transition point and embryonic genome activation occur. Blastocyst formation is essential for further development including implantation and pregnancy. It is important to evaluate embryo quality before transfer; however, we know little about the preimplantation stage embryo apart from gross morphological descriptions, cleavage timing and the ploidy status of selected chromosomes. Current studies on cellular composition and gene expression profiles in primate blastocysts require invasive techniques, but are critical to our long-term objectives of understanding and improving embryonic development. Such studies in the human are impacted upon by the scarcity of embryos available for research and by associated ethical concerns. However, because of the close physiological and genetic similarities between primates, the rhesus monkey is an excellent animal model for biomedical research on primate embryonic development. This animal model can also be used as an important experimental tool in the development and evaluation of novel technologies such as embryonic stem (ES) cell-based therapy. We believe that, in this case, translational research in a monkey model is absolutely essential before human clinical use, not only to demonstrate therapeutic efficacy, but also to allay safety concerns.

Oct4 expression in mammalian blastocysts Although molecular mechanisms underlying the separation of ICM and trophectoderm cell lineages are still poorly understood, it is known that the transcription factor Oct4 is essential for ICM development. Oct4, also named Oct3, belongs to the Pou (Pit, Oct, Unc) family of domain transcription factors. Expression of Oct4 is detectable throughout oogenesis and preimplantation development. However, in the mouse, Oct4 expression at the blastocyst stage is restricted to the ICM only, and later to the early epiblast, and finally confined to the developing germ cells6,9,10. The role of Oct4 has been elegantly demonstrated in mutant mice. Oct4 mutant mouse embryos develop into blastocyst-like structures containing trophectoderm but no ICM. Thus, Oct4 is essential for ICM

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development and has been used as a key marker for pluripotent cells such as embryonic stem (ES) or embryonic germ (EG) cells. In the mouse, the two cell populations responsible for the ICM and trophectoderm can be identified at the morula stage. According to the inside-outside hypothesis, the inner and outer cell layers in the morula develop into ICM and trophectoderm, respectively11. Conversely, in porcine and bovine embryos, compaction and cell allocation into ICM and trophectoderm seems to be an independent and random process12. Mouse embryos form an egg cylinder stage, and hatching and implantation occur almost simultaneously, whereas in farm animals blastulation is followed by the germinal disc stage and implantation is delayed following hatching13,14. These differences in the timing of genetic and morphological transitions may relate to differences in Oct4 expression. The protein produced by Oct4 expression has been detected in the ICM and the trophectoderm of cattle and porcine expanded blastocysts. We have been particularly interested in the pattern of Oct4 expression in primate embryos. Immunohistochemical examination of whole-mounted expanded rhesus monkey blastocysts reveals expression of Oct4 in the ICM as well as in trophectoderm. Oct4 protein is present in the nuclei of ICM cells, whereas a diffuse distribution of the signal, primarily in the cytoplasm of trophectoderm cells, is observed (Figure 12.2). Hatched blastocysts, however, show strong Oct4 expression in the ICM, with no detectable signal in the TE (Figure 12.2(d)). An Oct4 expression level was also studied in human blastocysts by reverse transcriptase-polymerase chain reaction (RT-PCR), and the mean Oct4 expression was 31 times higher in the ICM than in trophectoderm10. Interestingly, Oct4 was not detected in an ‘empty blastocyst’ lacking an ICM. This may suggest that Oct4 plays a similar role in maintaining pluripotency in humans and nonhuman primates as it does in mice. Mouse, human and monkey ES cells also express Oct4, and differentiation of these pluripotent cells is associated with loss of Oct4 signal. Thus, Oct4 is a candidate regulator in pluripotent and germ-line cells and is essential for the initial formation of a pluripotent founder population in the mammalian embryo. From a practical perspective, Oct4 expression can be used as a suitable marker to establish and maintain primate ES cell lines.

Cloning by nuclear transfer and embryo twinning In the past decade, tremendous progress has been made in the development of nuclear transfer (NT) technology. Live offspring have been produced in sheep, cattle, mouse, goat, pig and rabbit by nuclear transfer from differentiated somatic cells15–22. The possibility of cloning non-human mammals from somatic cells carries potential applications in a basic research context as well as at the applied level. Understanding the mechanisms of nuclear reprogramming after somatic cell nuclear transfer not only could lead to improving the efficiency of routine NT protocols but also could enhance our knowledge of early mammalian development. Improved efficiencies in producing healthy offspring by somatic cell nuclear transfer will allow its commercial use in agriculture and pharmaceutics. Coupling gene targeting with NT technology extends the possibility of producing transgenic and knock-out animals in species where it has not heretofore been possible23–25.

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The first requirement for cloning protocols is a reliable source of high-quality oocytes matured in vitro or in vivo following ovarian stimulation. Mature metaphase II (MII) oocytes are then enucleated using micromanipulation (Figure 12.3). Confirmation of metapbase spindle removal by staining with DNA-specific fluorochromes and epifluorescence is important, since artificial activation may induce parthenogenetic development of non-enucleated oocytes and developmental chaos after NT. The absence of a nuclear membrane makes it almost impossible to visualize the MII oocyte’s spindle under conventional microscopy. Blind enucleation based on removal of the first polar body and adjacent cytoplasm may ensure a high degree of complete enucleation in freshly matured MII oocytes. However, within 2–3 h of first polar body extrusion, polar body movement within the perivitelline space makes blind enucleation impractical and inefficient. The next step in NT is introducing the donor DNA (nucleus), from cultured or freshly isolated cells, into the enucleated oocyte (cytoplast). Conventionally, this is achieved by placing an isolated intact donor cell into the perivitelline space of the cytoplast followed by electrofusion. Alternatively, an isolated nucleus is injected directly into the cytoplast, similar to sperm transfer in the ICSI procedure (Figure 12.4). Synchrony between the nuclear donor cell and the cytoplast is important for maintaining the correct ploidy in the resulting reconstructed embryo. When a donor nucleus is transferred into an MII cytoplast prior to activation, high levels of maturation/meiosis/mitosis-promoting factor (MPF) induce rapid donor nucleus envelope-breakdown followed by premature chromosome condensation (Figure 12.5). This is the ‘normal’ response observed in many species, and may presage the reprogramming process necessary for successful full-term development. Of course, somatic cell nuclear transfer protocols bypass sperm-induced oocyte activation. Instead, artificial activation strategies designed to initiate cleavage and development are employed. The prerequisite oscillation in intracellular calcium concentrations followed by MPF inactivation can be accomplished, more or less, by exposure to ionophores and general protein synthesis or phosphorylation inhibitors. Cytoplast activation is followed by pronuclear formation and cleavage of the reconstructed embryo (Figure 12.6). For normal full term development of embryos created by NT from somatic cells, genes normally expressed during embryogenesis, but silent in the somatic donor cell, must be reactivated in an appropriate temporal and spatial manner. The genetic reprogramming required to reverse many, if not all, of these epigenetic changes in the somatic nucleus after NT is radically different from the process that occurs during natural gametogenesis, and must take place within the short interval between NT and the time of embryonic genome activation. While the production of NT embryos is now relatively routine in several species, the establishment of viable pregnancies and live offspring following transfer into recipients remains challenging, owing to high fetal and neonatal losses. These low success rates most likely reflect poor or incomplete genetic reprogramming of the donor nucleus followed by improper gene expression. DNA methylation is one of the epigenetic modifications controlling gene expression, and global methylation patterns are dynamic during preimplantation development and dramatically different from those seen in somatic nuclei. In several mammalian species studied including mice, rats, pigs and cattle, preimplantation embryos undergo genome-wide demethylation after fertilization followed by remethylation26. Somatic cell NT embryos showed a slight reduction in

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methylation consistent with active demethylation, but demethylation was subnormal. De novo methylation, resembling the methylation patterns of somatic cells, occurred precociously in many NT embryos26. This indicates that low efficiencies in somatic cell NT may to some degree be due to improper reprogramming as reflected by methylation levels. The advantage of producing identical animals for biomedical research is that, in general, a substantial reduction in animal number requirements can be realized27. Additionally, genetically identical animals are absolute requirements for some experimentation, for instance when immune system function is under study. Somatic cell NT may eventually provide a solution, given marked improvements in fetal and neonatal outcomes, although true clones or animals that are 100% genetically identical will not result, because of differences in the source of nuclear and mitochondrial DNA. The production of monozygotic offspring by manipulation of the preimplantation embryo remains an important alternative. Two approaches have been established in several mammalian species: blastomere separation at cleavage stages and blastocyst bisection. The ability of isolated blastomeres, either singly or in pairs, from 2-, 4- and 8-cell stage embryos to support term pregnancies and to produce genetically identical animals has been described in the mouse28, rat29, goat30, horse31 and, on repeated occasions, in cattle32,33. Although the usual outcome in producing monozygotic offspring is a singleton pregnancy, twins, triplets and even quadruplets have been reported upon transfer of onequarter embryos in cattle34,35. On average, 25–40% of live offspring resulted after transfer of in vivo-produced and -twinned cattle and sheep embryos as identical sets. However, in the mouse, rabbit and pig, the incidence of monozygotic twinning among offspring reached only 2–5%36. Forblastomere separation, zonae pellucidae of 2–4-cell stage embryos are enzymatically removed and then individual (from 2-cell embryos) or paired (from 4-cell embryos) blastomeres are separated mechanically aspirated into a micropipette and transferred into surrogate zonae pellucidae immobilized on a holding pipette (Figure 12.7). A convenient measure of the resulting demi-embryo viability is growth to the blastocyst stage in vitro compared with intact, or IVF- or ICSI-produced controls. The splitting of uterine stage embryos (morulae/blastocysts) has also led to the production of monozygotic offspring. Embryos can be recovered non-surgically by flushing the uterus of mated animals or by application of the assisted reproductive technologies. For bisection, the blastocyst is immobilized with a holding pipette, positioned across from the ICM, in a micromanipulation chamber on the stage of an inverted microscope. A surgical microblade attached to a micromanipulator is used to split the embryo, with even distribution of ICM and trophectoderm into each demiembryo (Figure 12.8). The zone-free demi-embryos produced by bisection are placed in culture and monitored for re-expansion. The number of identical animals that can be produced by blastomere separation or blastocyst bisection is realistically limited to twins or triplets, as the developmental potential of one-quarter embryos and less is poor, secondary to inadequate cell numbers to support the allocation of cells required during blastulation, as evidenced by extensive experience in agricultural species37. Thus, there may be insufficient embryonic mass for normal development or for the appropriate signalling that must occur between embryo and host. Demi-embryos separated at the 2- or 4-cell stage grow to the blastocyst stage at

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the same rate as intact controls, and contain, as evidenced by differential staining and confocal microscopy, half the number of cells38. Demi-blastocysts, when compared as averaged values and independent of the twinning method, also maintain allocations of ICM and trophectoderm cells with ICM/trophectoderm or ICM/total cell ratios the same as those of intact controls. Our preliminary results38 suggest that, for specific biomedical and genetic purposes, pairs of monozygotic monkeys can be produced by either embryo twinning technique, although we have yet to realize a monozygotic twin pregnancy to term. While spontaneous monozygotic twinning has been reported following blastocyst transfer in women39, it seems unlikely that the twinning approaches described here in monkeys will be used clinically at least in the near future.

Blastocyst evaluation by differential staining As noted above, the embryo at the blastocyst stage comprises a morphologically distinct ICM and trophectoderm. Clearly, both of these cellular types are required for normal implantation and pregnancy to ensue. The possibility that differences in cell allocation and count could impact upon viability has led to the development of specialized techniques for quantifying these cells. Such examinations are most commonly performed by differential staining (see reference 40 for review). One procedure is based on partial lysis of trophectodermal cells with complement, following labelling with species-specific antibody41. The ICM, however, is protected from antibody exposure by the selective permeability of the outer trophoblast layer. The embryo is stained with the DNA-specific dye, propidium iodide, which does not penetrate intact membranes and stains only partially lysed trophectodermal cell nuclei. The embryo is then counter-stained with a different DNA-specific dye, bisbenzimide (Hoechst 33342), which penetrates the membranes and labels the ICM. A modification of this technique has been developed that avoids the need for speciesspecific antibodies42. In this case, trophoblastic cell surface proteins are first labelled with trinitrobenzenesulfonic acid (TNBS) and subsequently recognized by a universal TNBSspecific antibody. The advantage of this approach is that all reagents are commercially available and suitable for any species. In yet another approach, the trophectodermal layer can be permeabilized by treatment with calcium ionophore combined with propidium iodide43. Calcium ionophore treatment triggers an osmotic response and formation of cell membrane vesicles, leadingto permeation of trophectoderm cells exclusively. The ICM cells remain intact, and can be differentially counterstained with bisbenzimide. The final examination of whole mounted specimens is carried out under a fluorescence microscope, equipped with excitation and emission filters appropriate for the fluorochromes in use. ICM nuclei labelled with bisbenzimide appear blue, while trophectoderm nuclei labelled with both bisbenzimide and propidium iodide appear pink or red (Figure 12.9). The numbers of ICM and trophectoderm nuclei can be counted directly under the microscope. More precise imaging can be performed using confocal microscopy, where different lasers can be used to excite bisbenzimide and propidium iodide. Fluorescent light in the specific spectral range is detected simultaneously in separate channels for either the propidium iodide or the Hoechst stain. Following this,

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optical sections approximately 0.5 µm in thickness can be sampled at 1–5-µm intervals throughout the entire blastocyst. Digital optical sections are analyzed by combining them into three-dimensional images using image-processing software such as MetaMorph 4.5. This approach allows automatic or semi-automatic cell counts of each section, and minimizes errors. In experimental embryology, differential staining techniques have numerous applications, such as defining the ‘normal’ allocation of ICM and trophectoderm cells in blastocysts and then comparing allocations for in vivo-versus in vitro-produced blastocysts. The impact of different culture media or embryo manipulations (blastomere biopsy, twinning, NT) on development can also be assessed. In the clinical arena, the technique could help correlate cell numbers and allocations with morphological criteria of blastocyst quality. This approach would ultimately improve the non-invasive evaluation and selection of blastocysts for intrauterine transfer or cryopreservation.

Primate embryonic stem cells: pluripotency assessment Embryonic stem (ES) cells and embryonic germ (EG) cells are pluripotent cell lines as derived initially from mouse preimplantation embryos44,45 and primordial germ cells46,47. Application of mouse ES cell technology coupled with gene targeting for the production of mice carrying predetermined genetic alterations has become a revolutionary tool for the study of mammalian gene function (for review see reference 48). ES and EG cells are morphologically distinct, and express specific cellular markers. They remain immortal and undifferentiated under certain conditions, and their pluripotency has been demonstrated both in vitro and in vivo. In the absence of feeder layers in vitro, ES cells are capable of differentiating into a variety of cell types representing all three embryonic germ layers. In vivo, when injected into immunocompromised mice, ES cells can form teratomas also containing cells representative of all three germ layers, however, this property is shared with mouse and rat visceral endoderm (yolk sac)49 and, therefore, may not be the most definitive measure of pluripotency. The ultimate measure perhaps, despite feasibility limitations, is the participation of ES cells in chimeric animals. Combined with host cells in a chimeric embryo, ES cells in the mouse can participate in development of all adult tissues, including germ cells50. Human pluripotent (ES and EG) cells have been isolated recently both from ICM cells of IVF-produced blastocysts51 and from primordial germ cells recovered from gonadal ridges and mesenteries of 5–9-week-old fetuses52. The use of such cells may eventually revolutionize the practice of medicine, with implications as far-ranging as our understanding of human embryogenesis and the development of transplantation therapies. The potency of human ES cells is important to establish as a prerequisite to clinical research that evaluates human ES cell use for cell, tissue or organ replacement/repair purposes. Pluripotency can most appropriately be established by quantitating participation of ES cell derivatives in chimeric embryos and in all cell lineages of chimeric fetuses and term infants, an undertaking that cannot be done in the human for ethical reasons. However, because of the close similarities between human and monkey ES cells, the use of rhesus monkey ES cells would serve as a valuable biomedical research model, and should catalyze progress towards clinical applications of ES cells.

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Rhesus ES cells proliferate indefinitely in vitro as undifferentiated stem cells, and retain the potential to differentiate into derivatives of trophectoderm and all three embryonic germ layers in vitro or in vivo after injection into immunodeficient mouse53,54. Based on morphology and the presence of specific markers (alkaline phosphatase activity, Oct4, cell-surface markers stage-specific embryonic antigen-3 (SSEA-3), SSEA4, TRA-1–60 and TRA-1–81) (the high-molecular-weight glycoproteins), rhesus ES cells resemble early totipotent embryonic and human embryonic carcinoma (EC) cells54. Presumptive ES cell lines with in vitro pluripotent characteristics similar to mouse ES cells have been established in several mammalian species, but only a few of them are able to generate chimeras: rat55, rabbit56, pig57 and cow58. Although the precise requirements for maintaining pluripotency in such cell lines are unknown, the rigorous application of specialized culturing techniques as well as the use of early passage cells significantly increases the chances of obtaining highly chimeric animals. The methods of production of ES cell-embryo chimeras currently fall into three groups: microinjection, aggregation and coculture. The predominant method used so far has been injection of 10–15 ES cells into the blastocoel cavity of blastocysts, followed by embryo transfer into pseudopregnant recipients50. Simplified techniques have been developed, based on the ability of ES cells to aggregate readily with cleavage stage embryos. For example, ES cell clumps aggregated with 4–8-cell stage zona pellucida-free mouse embryos or were sandwiched between two tetraploid embryos59. The use of tetraploid embryos is predicated on the observation that tetraploid host embryonic cells are effectively selected against in the developing chimeric embryo, producing 100% ES cell-derived mice; the tetraploid cells are well represented in the extraembryonic membranes and tissues. The third alternative employs simple short-term coculture of 8cell stage, zona-free embryos on a ‘lawn’ of disaggregated ES cells60 with the chimeric embryos cultured overnight to the blastocyst stage before transfer into recipients. The aggregation and coculture approaches involving zona-free embryos may not be an option for chimera production in mammalian species that require the presence of a zona pellucida during preimplantation development. A variation on the blastocyst injection and aggregation technique, which seems to be more efficient, involves injection of ES cells into cleavage stage embryos, where 10–15 ES cells are injected into the perivitelline space61–64. A similar approach has been successfully used to generate bovine ES cellderived chimeras after injecting ES-like cells into 8-cell stage embryos58. Injection of ES cells into 4–8-cell stage host embryos has been shown to be more efficient than blastocyst injection in terms of ES cell contribution to somatic and germ line tissue in the resultant chimeras. This approach also carries the possibility of confirming ES cell participation in chimeric blastocysts in vitro before further in vivo studies are undertaken. While phenotypic markers are readily available in mice, cellular markers are also important in assessing the extent of ES colonization of chimeric blastocysts. The lac-Z. reporter gene, coding for β-galactosidase whose activity can be followed in situ directly on whole-mounted embryos, has been used for this purpose in the mouse61,63. The presence of bovine ES-like cell derivatives, transgenic for lac-Z in multiple tissues of 5month-old chimeric animals, was identified and quantified by Southern blot analysis and fluorescence in situ hybridization (FISH) owing to the presence of the lac-Z reporter gene58. Thus, transfection of rhesus monkey ES cells with a lac-Z. or green fluorescence protein (GFP) construct is of considerable importance to success in the identification of

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ES cell derivatives in chimeric fetuses and offspring. Unfortunately, we have not yet succeeded in producing GFP-positive monkey ES cells. In an alternative approach, PKH26, a red fluorescent cell linker was used, based on its established application in cell labelling and short-term tracking. PKH-26 labelling has also shown low toxicity, with no cross-contamination of the label to neighboring cells. Rhesus monkey ES cells were dispersed into single cells or small clusters of 2–3 cells each and labelled with PKH-26. A total of 10–15 labelled cells were then injected into embryos at the 4–8-cell stage (Figure 12.10a and b), and the incorporation of cells into the chimeric embryos was monitored under epifluorescent microscopy throughout preimplantation development. Monitoring labelled, injected cells provides valuable insights into the proliferative activities and localization of ES cells in morulae and blastocysts. Images taken from live embryos (Figure 12.10) clearly demonstrate ES cell colonization of both the ICM and trophectoderm. These chimeric blastocysts can be further analyzed by differential staining, followed by confocal microscopy and quantitation of ES derivatives in the ICM and trophectoderm. In summary, clear evidence of ES cell proliferation in early preimplantation development was obtained in monkey embryos with distribution into both the trophectoderm and ICM. These results demonstrate that chimeric embryos can be produced readily in the rhesus monkey by ES cell injection, and set the stage for potency determination in chimeric fetuses or offspring when permanently labelled ES cells become available.

Preimplantation genetic diagnosis Preimplantation genetic diagnosis (PGD) allows embryos to be screened for genetic disorders, followed by the selection of unaffected embryos for transfer before the establishment of pregnancy. Basically, the procedure of PGD involves two major steps, collecting diagnostic material via polar body or embryo biopsy and genetic testing for a specific genetic disorder by molecular or molecular-cytogenetic approaches. Couples either affected by or carriers of a specific genetic disorder can create embryos by IVF and have one or two blastomeres aspirated from the 6–8-cell stage embryo, or polar bodies can be removed from the oocyte or zygote. In the latter case, genetic analysis can detect a disorder if it is of maternal origin. The collected materials are then processed for genetic diagnosis. There are two major methods currently used for PGD testing, polymerase chain reaction (PCR) and FISH. PCR-based methodologies are mainly used to diagnose single-gene disorders, while FISH-based methodologies are used to detect numerical and structural chromosomal abnormalities. Although PGD has now been practiced for more than 10 years, there are still few centers worldwide that can effectively offer these clinical services to patients. As there are several cases of misdiagnosis reported65,66, concerns of accuracy along with substantial technical demands impact upon the widespread application of PGD. Misdiagnosis in PGD may reflect biological or technical factors. Biological factors indicate abnormalities inherent to the embryo(s) or blastomere(s) taken for genetic analysis, such as chromosome mosaicisms or nuclear abnormalities. The high rate of chromosome mosaicism detected in cleavage stage embryos67–69 has been regarded as a

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potential problem for the accuracy of single-cell PCR diagnosis for single gene defects or FISH diagnosis for chromosomal abnormalities or gender selection70,71. Technical factors include improper processing of diagnostic materials, contamination by exogenous genetic materials, amplification failure or inaccurate signal detection. As described above, many factors impact upon PGD when single cells are used; however, improved protocols will undoubtedly further reduce the misdiagnosis rate due to technical limitations72. Errors secondary to genetic diagnostic attempts involving only one or two cells remain. The biopsy of trophectoderm cells of blastocysts may provide a way of obtaining more diagnostic material for PGD testing, as 2–30 cells can be obtained from human blastocysts with various biopsy strategies73–75. Blastocyst biopsy has several potential advantages over cleavage stage or polar body biopsy. First, since blastocyst biopsy involves removal of cells from the trophectoderm, an exclusively extraembryonic lineage, the ICM, orcells that contribute to the embryo proper, should remain unperturbed. Second, as noted above, many more cells can be removed from the trophectoderm, providing the basis for more reliable genetic testing. Third, embryos that reach the blastocyst stage in culture may have an increased implantation potential, ensuring a higher overall success rate for PGD, when carried out at this stage. The feasibility of trophectoderm biopsy has been demonstrated in various species (rabbit76, mouse77,78, cattle79, marmoset80, human73) with several biopsy strategies including: trophectoderm incision, where a microblade inserted through a small hole made in the zona pellucida is used to excise trophectoderm cells; and zona drilling and excision, drilling with acidified Tyrode’s solution (Figure 12.11a) away from the ICM and dissecting trophectoderm after partial hatching with a microblade (Figure 12.11b and c). A laser-mediated approach has been used to perform zona drilling and subsequent trophectoderm biopsy on human blastocysts75. Development of biopsied blastocysts seems unimpaired, as it has been shown that more than 40% of manipulated human blastocysts reach the hatching stage after biopsy73. In addition, birth of normal offspring after trophectoderm biopsy has been reported in the mouse81 and marmoset80. Although trophectoderm biopsy has been performed on human blastocysts for research purposes for many years, this technique has not been widely applied in clinical PGD. A main concern has been the inefficient development of embryos to the blastocyst stage. Furthermore, only limited data are available in the human on the effects of trophectoderm biopsy on implantation, pregnancy rate and postimplantation embryonic development. With improvements in culture conditions, high blastocyst rates are now routinely achieved in vitro82,83. This should encourage further interest in blastocyst biopsy; however, a greater understanding of implantation and postimplantation development of biopsied blastocysts is still needed. This knowledge base should ideally be preceded by safety and efficiency assessments in animal models such as the monkey. In conclusion, the potential applications of PGD expand as our knowledge of the human genome increases. This technology will significantly impact upon the future of human assisted reproductive technologies, and could lead to use in fertile patients at genetic risk. Additionally, microarray technologies will allow the determination of gene expression patterns or the ability to analyze the genetic polymorphism within individual embryos, and, therefore, provide an opportunity to identify embryonic viability markers or specific genetic disorders. Clearly, these objectives provide incentives to overcome the

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current limitations of PDG, namely the limited genetic material available for diagnosis. Translational research in animal models, including the non-human primate, we believe will contribute significantly to the further development of PGD.

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65. Geraedts J, Handyside A, Harper J, et al. European Society of Human Reproduction and Embryology Preimplantation Genetic Diagnosis Consortium. Preliminary assessment of data from January 1997 to September 1998. ESHRE PGD Consortium Steering Committee. Hum Reprod 1999; 14:3138–48 66. Lewis CM, Pinel T, Whittaker JC, Handyside AH. Controlling misdiagnosis errors in preimplantation genetic diagnosis: a comprehensive model encompassing extrinsic and intrinsic sources of error. Hum Reprod 2001; 16:43–50 67. Munne S, Sultan KM, Weier HU, Grifo JA, Cohen J, Rosenwaks Z. Assessment of numeric abnormalities of X, Y, 18, and 16 chromosomes in preimplantation human embryos before transfer. Am J Obstet Gynecol 1995; 172:1191–9, discussion 1199–201 68. Delhanty JD, Harper JC, Ao A, Handyside AH, Winston RM. Multicolor FISH detects frequent chromosomal mosaicism and chaotic division in normal preimplantation embryos from fertile patients. Hum Genet 1997; 99:755–60 69. Kuo HC, Ogilvie CM, Handyside AH. Chromosomal mosaicism in cleavage-stage human embryos and the accuracy of single-cell genetic analysis. J Assist Reprod Genet 1998; 15:276– 80 70. Harper JC, Coonen E, Ramaekers FC, et al. Identification of the sex of human preimplantation embryos in two hours using an improved spreading method and fluorescent in-situ hybridization (FISH) using directly labelled probes. Hum Reprod 1994; 9:721–4 71. Delhanty JD, Wells D, Harper JC. Genetic diagnosis before implantation. Br Med J 1997; 315:828–9 72. Munne S, Marquez C, Magli C, Morton P, Morrison L. Scoring criteria for preimplantation genetic diagnosis of numerical abnormalities for chromosomes X, Y, 13, 16, 18 and 21. Mol Hum Reprod 1998.4:863–70 73. Dokras A, Sargent IL, Ross C, Gardner RL, Barlow DH. Trophectoderm biopsy in human blastocysts. Hum Reprod 1990; 5:821–5 74. Muggleton-Harris AL, Braude PR. Preimplantation diagnosis of genetic disease. Curr Opin Obstet Gynecol 1993; 5:600–5 75. Veiga A, Sandalinas M, Benkhalifa M, et al. Laser blastocyst biopsy for preimplantation diagnosis in the human. Zygote 1997; 5:351–4 76. Gardner RL, Edwards RG. Control of the sex ratio at full term in the rabbit by transferring sexed blastocysts. Nature (London) 1968; 218:346–8 77. Gardner RL. Manipulation on the blastocyst. Adv Biosci 1971; 6:279–96 78. Monk M, Muggleton-Harris AL, Rawlings E, Whittingham DG. Pre-implantation diagnosis of HPRT-deficient male and carrier female mouse embryos by trophectoderm biopsy. Hum Reprod 1988; 3:377–81 79. Betteridge KJ, Hare WCD, Singh EL. Approaches to sex selection in farm animals. In Brackett BG, Seidel GE, Seidel SM, eds. New Technologies in Animal Breeding. New York: Academic Press, 1981:109–25 80. Summers PM, Campbell JM, Miller MW. Normal in-vivo development of marmoset monkey embryos after trophectoderm biopsy. Hum Reprod 1988; 3:389–93 81. Gentry WL, Critser ES. Growth of mouse pups derived from biopsied blastocysts. Obstet Gynecol 1995; 85:1003–6 82. Gardner DK. Development of serum-free media for the culture and transfer of human blastocysts. Hum Reprod 1998; 13(Suppl4):218–25 83. Gardner DK. Schoolcraft WB. Culture and transfer of human blastocysts. Curr Opin Obstet Gynecol 1999; 11:307–11

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Figure 12.1 Rhesus monkey preimplantation embryo development in vitro. (a) Pronuclear stage zygote

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with male and female pronuclei, 12 h after fertilization by intracytoplasmic sperm injection (ICSI); (b) day-1, 2cell stage embryo (day of fertilization=day zero); (c) day2, 4-cell stage embryo; (d) day-3, 8-cell stage embryo; (e) day-4, morula stage embryo; (f) day-5, compact morula. Note that individual blastomeres have maximized their intracellular contacts and compacted into a tight cell mass; (g) day-7 or -8, expanded blastocyst with a single flat layer of trophectoderm surrounding a fluidfilled blastocoel and the inner cell mass

Figure 12.2 Oct4 expression in rhesus monkey blastocysts. (a) Epifluorescent microscopy of a day-8 rhesus monkey

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expanded blastocyst stained with the DNA-specific dye, DAPI. (b) The same blastocyst labeled with Oct4 antibody and secondary antibody conjugated with Cy3. Note the distribution of the immunofluorescence signal in both the ICM and trophectoderm. (c) More advanced day-9 hatched blastocyst. (d) Note the strong expression of Oct4 in the ICM, with no detectable signal in the TE

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Figure 12.3 Nuclear transfer: preparation of the cytoplast. Enucleation of a metaphase II (MII), rhesus monkey oocyte. (a)

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Epifluorescence visualization of the metaphase spindle labeled with bisbenzimide. In the freshly matured MII oocyte, the spindle is localized in the cytoplasm/cortex beneath the first polar body. (b) Removal of the first polar body and the metaphase spindle by aspiration into an enucleation pipette. (c) Enucleated oocyte (cytoplast) under Hoffman optics

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Figure 12.4 Nuclear transfer: creation of the fusion pair. Introducing the donor DNA material into the cytoplast. (a, b) Placing the intact donor cell into

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the perivitelline space with a micropipette. This procedure is followed by electrofusion that induces membrane fusion and incorporation of the entire nuclear donor cell. (c) Direct injection of an isolated nucleus into the cytoplast, similar to sperm transfer in the intracytoplasmic sperm injection (ICSI) procedure, is another method of introducing the donor nucleus.

Figure 12.5 Nuclear transfer: cytoplast activation. Premature chromosome condensation. High levels of maturation/meiosis/mitosis-promoting factor (MPF) of the non-activated cytoplast induce rapid nuclear envelope breakdown in the donor nucleus followed by premature chromosome condensation (PCC). Activation is subsequently achieved by exposure to ionophores and general protein synthesis or phosphorylation inhibitors

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Figure 12.6 Nuclear transfer: embryo development. (a) Pronuclear-stage, rhesus monkey, somatic cell nuclear transfer (NT) embryo 10 h postactivation with a single pronucleus containing multiple nucleoli. (b) Rhesus monkey, somatic cell, NT embryos at the 4–6-cell stage. Note the presence of a clear distinctive nucleus in each blastomere

Figure 12.7 Embryo twinning by blastomere separation. (a) 2-cell stage rhesus monkey embryos after removal

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of zonae pellucidae; (b) a single blastomere from a 2-cell embryo in the transfer pipette, positioned for placement inside a surrogate zona pellucida which is immobilized on a holding pipette; (c) monozygotic, 1cell, demi-embryos produced by blastomere separation; (d) hatching, monozygotic, demi-blastocysts after in vitro culture for 8 days

Figure 12.8 Embryo twinning by blastocyst bisection. (a) Expanded rhesus monkey blastocyst immobilized with a holding pipette positioned opposite the inner cell mass (ICM); (b) initial bisection of the ICM with a surgical microblade while holding the blastocyst; (c) completion of the

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bisection process after release from the holding pipette; (d) monozygotic, demi-blastocyst immediately upon completion of the bisection step

Figure 12.9 Differential staining of a rhesus monkey blastocyst. Inner cell mass (ICM) nuclei labeled with bisbenzimide appear green, while trophectoderm nuclei labeled with both bisbenzimide and propidium iodide appear pink or red. Individual cells can be quantitated by this technique

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Figure 12.10 Production of rhesus monkey chimeric embryos with embryonic stem (ES) cells. (a) Injection of a total of 10–15 ES cells into the perivitelline space of a 4–8cell stage embryo (Hoffman optics); (b) epifluorescence microscopy of embryos injected with PKH-26 labeled ES cells; (c) injected embryo cultured to the hatching blastocyst stage; (d) the same blastocyst under epifluorescence microscopy showing colonization of labeled ES cell derivatives in both the inner cell mass (ICM) and trophectoderm. The intensely fluorescent area is thought to arise from degenerating ES cells; (e) epifluorescent microscopy of a day-8, expanded rhesus monkey blastocyst stained with bisbenzimide (blue); (f) the same embryo displaying labeled ES cell contributions (red) to the ICM and trophectoderm

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Figure 12.11 Trophectoderm biopsy for preimplantation genetic diagnosis. (a) Expanded rhesus monkey blastocyst immobilized with a holding pipette. The zona drilling pipette filled with acidified Tyrode’s solution is positioned opposite the inner cell mass (ICM); (b) hatching of trophectodermal cells after zona drilling and culture for several hours while the ICM remains inside the zona.

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The extent of hatching can be regulated by the culture interval; (c) initial excision of the hatched trophectodermal cells with a surgical microblade while holding the blastocyst; (d) embryo immediately upon completion of the biopsy step. Note that biopsy did not cause collapse of the blastocyst, indicating minimum damage; (e) biopsy material containing 15–20 trophectodermal cells

13 The moral status of the human blastocyst Howard W.Jones, Jr

As interest in the human blastocyst waxes not only as a means for improving clinical pregnancy rates with in vitro fertilization (IVF) but also in the basic science arena as a source of embryonic stem cells, it is fitting that the ethical/moral status of the blastocyst be examined once more. Here, as an aside, it is perhaps appropriate to comment on the difference between ethics and morality. This was discussed in a chapter by Ernlé Young in The Human Embryonic Stem Cell Debate1. Young holds that ethics analyzes an action or situation from the point of view of natural reason, whereas morality analyzes an action or situation from the point of view of traditional canon law. Different views on the moral status of the preembryo (of which the blastocyst is a part) became public issue when IVF was first introduced as a treatment modality to assist in overcoming infertility2. At the heart of the matter is determining when, during development, personhood is achieved. Personhood in this context means the achievement of a developmental status that deserves protection by society. From a clinical view this is not important for the blastocyst which, when transferred, implants and develops or which, when transferred, fails to develop, but it is important for those that are not transferred. From the point of view of basic scientists and embryologists, great importance is placed on the moral/ethical status of blastocysts not transferred or those used for experimentation. The acquisition of personhood or ensoulment has been considered by at least three disciplines of our culture. Note that ensoulment is equated with personhood in this context. It is generally agreed that the soul and person are different entities, i.e. the soul is a somewhat ethereal entity with quite an individual identity, whereas the person can be considered a civil designation with certain social and legal rights and responsibilities. However, if after ensoulment there is the implication that civil protection is required, then the soul and the person are the same in this particular context.

Canon law according to the classical tradition, canon law according to the current tradition, Jewish tradition and law, Islamic tradition and law, American civil law, and natural reason Canon law: the classical tradition The classical tradition can be traced to Aristotle and perhaps even before, to Egyptian culture (Figure 13.1). In De Anima, Aristotle indicated a belief that the individual acquired three different souls in sequence (approximately 350BC). First came a vegetable soul, then an animal soul and finally at birth a rational soul. Aristotle specified a time

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sequence. For males, the animal soul was acquired at approximately 40 days of development, while females acquired theirs at closer to 80 days. This concept of multiple souls was adopted by the early church fathers, and is seen in the writings of St Thomas, St Augustine, St Jerome and many others. The monk and teacher, Gratian, then summarized the matter in his influential publication, Decretum (AD1140), a cornerstone text of canon law (Figure 13.2). He codified several revisions of law having to do with a variety of ecclesiastical matters, but particularly with regard to abortion. The summary therefore is very relevant and authoritative to the matter at hand. According to Gratian, abortion was not murder if the soul had not been infused. In other words, he accepted the notion that the soul had not been infused until some point during development after implantation. As proof, he offered the following: first, a statement by St Augustine regarding destruction of the non-animated fetus; second, the fact that the body must be formed to accept the soul, as for example in the case of Adam; and third, the statement of St Jerome saying that murder requires a formed fetus. There can be little doubt that the early church fathers, before and during the Middle Ages, clearly accepted the notion that ensoulment did not occur until some time during development. We can interpret this in modern terms by saying that the preembryo, and probably the embryo, were not considered by these early Christian leaders as being ensouled, i.e. they were not persons and thus did not deserve special protection by society. Canon law: the current tradition All Christians have been born and educated in a world in which the tradition of canon law of the Roman Church states that ensoulment occurs with fertilization. This has therefore been referred to as the current traditional concept within that church. This concept of personhood was powerfully underlined by Pope Pius IX who convened the XXth Ecumenical Council of the Roman Church, commonly referred to as Vatican I, which was in session from 1869 to 1870. The approval of the content of several of the popes’ encyclicals and other writings by Vatican I had the effect of establishing or modifying canon law, the basic laws of governance of the Roman Church. Our interest here centers on two laws promulgated by Vatican I. The first is Tastor Aeternus’, which declared that in matters of faith or morals the pope could speak with infallibility, and Apostolicus Sedis’, in which punishment is outlined for those who commit certain crimes. The highest punishment of excommunication is prescribed for perpetrators of several acts, including those seeking to procure (provide/bring about) abortion if the desired effect ensues. The significant aspect of Apostolicus Sedis’ is that it no longer recognized a period during embryonic development before which excommunication did not apply. This has generally been interpreted and often cited as the concept that resulted in the modification of canon law to mean that ensoulment, i.e. personhood, was acquired with fertilization. This matter has been thoroughly reviewed in The Crime of Abortion in Canon Law by Father John Huser3. Important to recognize is the distinction between church legislation, i.e. canon law, and church practice as annunciated by clergy in good standing. This is especially true at the pastoral level but at other levels as well. This view is clarified in Health and Medicine in the Catholic Tradition by Father Richard A.McCormick4.

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It is astonishing to realize that it was as recent as 1869 and after 18 centuries of the classical tradition that canon law concerning punishment for abortion was altered from the Aristotelian teaching, which was adopted by the early church fathers, to what we now regard as the traditional view of the Roman Catholic Church and several other religious traditions. Jewish tradition and law According to Jewish tradition and law, the unborn fetus is not considered a person until it has been born5. Up to 40 days after fertilization, the preembryo is considered as ‘mere fluid’ or ‘water’, and regarded as part of the mother’s body. These facts form the basis for the Jewish legal view on abortion, allowing it to be carried out under certain circumstances. Abortion on demand, however, is repulsive to the ethics of the Talmudic Halakha, although there are many situations for which a pregnancy might be terminated, including saving the mother’s life. Furthermore, the creation of a preembryo for research purposes might be allowed if there is a true opportunity for the sperm owner to benefit in having a child as a result of the research. Interestingly Jewish law forbids the destruction of a preembryo if it possesses the potential for implantation, but one already hatched from its zona pellucida is regarded as having lost this potential, thus becoming acceptable for use in approved research5. Islamic tradition and law It is relevant to note that the understanding of classical Greece can also be found in the Islamic tradition, as well as others, either stemming from Greek philosophy or borne of independent origin. These derivations are wonderfully summarized in ‘The human embryo: Aristotle and the Arabic and European traditions’6 by Father Gordon Dunstan, Professor Emeritus of Moral Social Theology at the University of London and research fellow at the University of Exeter, as well as by others. Dunstan stated the ‘Quran left us no doubt that the fetus undergoes a series of transformations before becoming human.’ Islamic views generally place ensoulment on or after the 120th day, after three 40-day periods of development7. The preembryo itself has no precisely defined moral status. According to a publication from the National Bioethics Advisory Commission, research on preembryos and stem cells is regarded as ‘an act of faith in the ultimate will of God as the Giver of all life, as long as such an intervention is undertaken with the purpose of improving human health’8. American civil law Generally speaking, American civil law has not recognized the early conceptus as a person entitled to rights associated with personhood. Before medical technology introduced the possibility of extracorporeal preembryos, American jurisprudents addressed rights-related issues pertaining to a conceptus or fetus largely in the context of procreative or abortion-related privacy rights of the adult seeking to create or not create a child. As a long line of United States Supreme Court cases made clear, the ‘law affords constitutional protection to personal decisions relating to marriage, procreation,

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contraception, family relationships, child rearing, and education’ (Planned Parenthood of Southeast Pennsylvania v. Casey (1992)). The Supreme Court ruling in Roe v. Wade clearly established the pre-eminent right of women to terminate a pregnancy up to the point of viability. At the same time, the Court acknowledged that the state had an ‘important and legitimate interest in protecting the potentiality of human life’ (1973). The status of preembryos has been the subject of only a handful of lawsuits, all civil suits involving the disposition those cryopreserved after social or marital circumstances changed for the adults involved. The three earliest cases were York v. Jones (1989), Davis v. Davis (1992), and Kass v. Kass (1998). Since then, a growing number of courts have been confronted with these issues. In York v. Jones, the case involved transporting a preembryo from one clinic to another. The court ultimately considered the preembryo to be personal property belonging to the parents, and refused rights of protectorship requested by the clinic performing the freezing of and caring for the preembryo during cryostorage. In the second case, Davis v. Davis, a divorcing couple was unable to agree whether cryopreserved preembryos should be given to the wife for future attempts at motherhood or to the husband for destruction because of his desire to avoid parenthood. In this case, thetrial court considered the preembryos as persons and awarded ‘custody’ to the wife. Nonetheless, the Supreme Court of Tennessee reversed the decision and squarely rejected any characterization of the preembryos as persons, either under state or federal law, and concluded that they must occupy an interim position which entitles them to another form of respect because of their potential for human life. Finally, in Kass v. Kass, a New York court concluded that a divorced couple’s previous wishes in this ‘quintessentially personal private decision’, wishes evidenced in a signed cryopreservation agreement, should rule. Although the courts differ somewhat both in the characterization of preembryos and over who should have the right to control them, no court has ultimately recognized the preembryo as a person or entity entitled to the full panoply of rights associated with personhood, or even to the more limited rights associated with fetal viability. Natural reason Natural reason accepts that biology is unable to identify a point in the development of an individual that signifies the acquisition of what we define as personhood. It furthermore rejects the external infusion of an element which can be defined as personhood. Personhood, therefore, according to natural reason, develops in a Darwinian sense, i.e. slowly and through biological development. This concept recognizes that, for practical purposes, it is necessary for society to set certain arbitrary times as to when a preembryo, embryo, fetus or living person may acquire various rights as provided under civil law. Underscoring this is the belief that an ideal society expresses the will of the people and does so in a democratically organized manner. This point of view is eloquently set forth in Darwin’s Dangerous Idea: Evolution and the Meanings of Life by Daniel Dennett9.

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Blastocysts and research Why is this important at this time? It has to do with stem cells. The public is concerned about, and wants to be informed about, the exact stage of development from which stem cells are obtained. The concern is that, if the removal of pluripotent stem cells results in the cessation of development, then that stage acquires significance in terms of whether or not it is beyond a trigger point that designates protection by society. To be sure, the processes of spermatogenesis, oogenesis, fertilization and development are a continuum. Nevertheless, there are milestones, i.e. clear marker events, which conveniently segment various stages of development, e.g. beginning of the embryonic period, beginning of the fetal period (Figure 13.3). The events of the first few days of development are so biologically unique that they deserve to be described and segregated. This is the preembryonic period previously described10. This turbulent period has the following characteristics: (1) Large numbers of abnormalities occur. Estimates vary as to the extent of these abnormalities which cause loss, but a conservative estimate is that at least two-thirds of the products of oocyte and sperm interaction are in some way defective, either chromosomally or at a molecular level. The carrier of these abnormalities is so abnormal that it never implants, or, if it does implant, it usually perishes very early in development. (2) During this early phase of development, most of the developing structures will be devoted to nourishing the subsequent embryo. The trophectoderm predominates and is the predecessor of the placenta and extraembryonic membranes that will be discarded at the time of birth. While the inner cell mass is recognizable during the blastocyst stage, it consists of only a few cells, rudiments of the actual embryo that will form. It is from these cells that stem cells may be obtained. (3) Before the actual embryo forms with its neural groove, twinning may occur. There is no guarantee that there will be a single individual until the end of the preembryonic period, i.e. until a single primitive streak develops (Figure 13.4). (4) An individual may not develop at all. As a result of fertilization, the products of fertilization may end up as a tumor, a hydatidiform mole or, even worse, a chorioepithelioma that may ultimately destroy the host. (5) The lack of specificity of individual development is illustrated by the fact that, during this interval, fusion of two preembryos can occur and result in the development of a single fetus. This is well established in the human where fusion occasionally takes place with preembryos of different sexes. The presumption must be that, if this is possible, there should be at least an equal number of instances where fusion occurs between two XX preembryos or between two XY preembryos. Thus, the primitive streak guarantees biological individuation and terminates the preembryonic period. Stem cells seem best obtained from the pluripotential cells of the inner cell mass of the blastocyst, a stage occurring during the preembryonic interval (Figure 13.5). For those who place the acquisition of personhood beyond this stage, using pluripotent cells from donated (and, otherwise, discarded) preembryos is carried out with a clear conscience. The recent discussions about stem cells through print, visual and audible media, and by both the general public and the scientific community, have indicated that there is considerable variation in the concepts of the stages of development and the significance thereof.

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The concept of a preembryonic stage occurring earlier than an embryonic one was introduced in 1986 by the American Fertility Society, now the American Society for Reproductive Medicine, through its Ethics Committee, with the issuance of the report of that committee11. At the same time, the term preembryo was also introduced independently by the Volunteer Licensing Authority, an arm of the Royal College of Obstetricians and Gynaecologists and the British Medical Research Council. The preembryonic stage was identified as the interval up to the appearance of the primitive streak, a stage which guarantees biological individualization at about 14 days. It is of some significance that, quite independently, the Ethics Advisory Board of the Department of Health, Education and Welfare in its report in 1979 designated the period up to 14 days as having attained special moral status. This period approved for research by certain ethical committees is not recognized under Jewish law5.

A final thought Surely there is controversy about almost every point in this discussion. This controversy whirls in the heads of patients, doctors, nurses and medical support staff, and in the heads of religious counsellors, ethicists, philosophers, teachers, legislators, lawyers, judges and all those who address these issues, including prospective parents. An examination of the roots of our belief systems may help in converting controversy to consensus. One thing seems clear: medical care-givers at all levels must understand why they believe what they do, so that they are capable of intelligent discussion of these issues with patients. Only through an ordered and rational thought process may we genuinely assist couples in becoming comfortable with the difficult decisions they may be forced to make in today’s new era of reproductive options.

References 1. Young E. Ethical issues: a secular perspective. In Holland S, Lebacqz K, Zoloth L, eds. The Human Embryonic Stem Cell Debate: Science, Ethics, and Public Policy. Basic Bioethics. Cambridge, MA: MIT Press, 2001; 163–74 2. Jones HW, Crockin SL. On assisted reproduction, religion, and civil law. Fertil Steril 2000; 73:447–52 3. Huser RJ. The Crime of Abortion in Canon Law: An Historical Synopsis and Commentary. Washington, DC: The Catholic University of America Press, 1942 4. McCormick RA. Health and Medicine in the Catholic Tradition: Tradition in Transition. New York: Crossroad, 1984 5. Schenker JG. Infertility evaluation and treatment according to Jewish law. Eur J Obstet Gynecol Reprod Biol 1997; 71:113–21 6. Dunstan GR, Seller MJ. The human embryo: Aristotle and the Arabic and European traditions. In Dunstan GR, ed. The Status of the Human Embryo: Perspectives from Moral Tradition. London: King Edward’s Hospital Fund for London, 1988:38 7. Wertz DC. Embryo and stem cell research in the United States: history and politics. Gene Ther 2002; 9:674–8

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8. Sachedina A. Islamic perspectives on research with human embryonic stem cells. In Ethical Issues in Human Stem Cell Research, Vol III, Religious Perspectives. Rockville, MD: National Bioethics Advisory Commission, US Government Printing Office, 2000:G1–6 9. Dennett DC. Darwin’s Dangerous Idea: Evolution and the Meanings of Life. New York: Simon & Schuster, 1995 10. Jones HW Jr, Schrader C. And just what is a preembryo? Fertil Steril 1989; 52:189–91 11. Ethics Committee of the American Fertility Society. Ethical considerations of the new reproductive technologies. Fertil Steril 1986; 46 (Suppl 1):1S–94S

Figure 13.1 Aristotle contemplating the bust of Homer (1653), by Rembrandt van Rijn. (New York Metropolitan Museum of Art)

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Figure 13.2 Gratian’s Decretum

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Figure 13.3 Eight-week fetus, 4 cm in length. Courtesy Lennart Nilsson, A Child is Born, 1990, Delacorte Press, p. 91. The early fetus is suspended in the amniotic fluid

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Figure 13.4 Neural tube, 2 mm in length. Courtesy Lennart Nilsson, A Child is Born, 1990, Delacorte Press, p. 77. The outer layer, the skin, is cleft by the groove of the neural tube. The swelling above is the rudimentary forebrain

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Figure 13.5 High magnification of the inner cell mass of a developing blastocyst. Photograph courtesy of L.Veeck

Glossary of terms Abembryonic Away from the embryo or away from the inner cell mass Acrosome reaction A reaction that occurs when a spermatozoon contacts the zona pellucida in mammals; the acrosome reaction involves a sequence of structural changes in the sperm acrosome, including the liberation of enzymes which are thought to assist sperm penetration through the zona pellucida Activation (oocyte activation) The process through which a secondary oocyte is stimulated to resume meiosis. This can occur by a penetrating spermatozoon or an artificial substitute Adherens junctions Junctions which provide strong mechanical attachments between adjacent cells, built from cadherins and catenins Adhesion molecules Surface ligands, usually glycoproteins, that mediate cell-to-cell adhesion. Their functions include the assembly and interconnection of various vertebrate systems, as well as maintenance of tissue integration, wound healing, morphogenic movements, cellular migrations and metastasis AHA or assisted hatching Assisted hatching; a procedure undertaken in in vitro fertilization (IVF) laboratories where a small hole is made in the zona pellucida of a preembryo to facilitate natural hatching Amnion The innermost membrane enclosing the embryo or fetus Aneuploidy Any deviation from an exact multiple of the haploid number of chromosomes, whether fewer or more Animal pole/vegetal pole In an oocyte, the animal pole represents the region from which the polar bodies arise. The vegetal pole is opposite the animal pole Apoptosis A form of cell death in which a programmed sequence of events leads to the elimination of cells without releasing harmful substances into the surrounding area. Apoptosis is also called programmed cell death or cell suicide. Apoptosis plays a crucial role in developing and maintaining health by eliminating old cells, unnecessary cells and unhealthy cells ART Assisted reproductive technology Aspermia Failure to form or produce an ejaculate Asthenozoospermia

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Production of an ejaculate with very poor motility Azoospermia Production of an ejaculate devoid of spermatozoa Biopsy A procedure where small pieces of tissue or individual cells are taken for microscopic study preimplantation genetic diagnosis, use in reproductive techniques or tissue culture Blastocoel The fluid-filled cavity of the blastocyst Blastocyst The mammalian conceptus in the postmorula stage. The cells of the blastocyst form a spherical shell enclosing the blastocoel, with one pole distinguished by the inner cell mass from which the embryo forms. The outer cell layer forms the first differentiated epithelial-like cell line, the trophectoderm Blastomere One of the cells produced by the cleavage of a fertilized oocyte; a cleavage cell Cadherins Membrane-bound proteins that mediate the binding of like tissues. Neural tissues express N-cadherins while surface ectoderm expresses E-cadherins Cavitation The process involving formation of the fluid-filled extracellular cavity within a compacted preembryo; this cavity increases in size to form the blastocoel Cell allocation The movement of cells to different regions or their assignment of different functions or purposes within an organism Centriole One of a pair of cellular organelles that are adjacent to the nucleus, function in the formation of the mitotic apparatus and consist of a cylinder with nine microtubules arranged peripherally in a circle Chorion The outermost fetal membrane that encloses the amnion, closest to the wall of the uterus Chromosome The structures in the cell that carry the genetic material (genes); the genetic messengers of inheritance. The human has 46 chromosomes, 23 coming from the oocyte and 23 coming from the spermatozoon Cleavage A series of cell divisions that occur early in development to divide the 1-cell zygote into a large number of blastomeres Cloning The production of identical copies of a living organism Coculture A laboratory technique that involves growing a specimen (in IVF, afertilized oocyte or preembryo) on a monolayer of feeder cells (e.g. epithelial cells) or medium that has been conditioned with such cells. These feeder cells are thought to enhance properties of the culture medium by absorbing toxins or releasing favorable growth factors

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Colchicine A chemical that disrupts microtubule formation Compaction A process through which a cleaving preembryo changes from a collection of individual cells into a solid mass with indistinguishable cell membranes. Compaction results from the formation of tight intercellular junctions which cause blastomeres to become closely apposed Conceptus The derivatives of a fertilized oocyte at any stage of development from fertilization to birth; includes extraembryonic membranes, as well as the preembryo, embryo or fetus. The products of conception; all structures that develop from the zygote, both embryonic and extraembryonic. A term commonly interchanged with the term preembryo during IVF treatment Corona radiata A closely apposed layer of follicle cells which surrounds the mature oocyte Corticosteroids Synthetic steroid hormones; may be used clinically in a prophylactic manner to suppress the immune response Crossing-over A process occurring during synapsis in which pairs of homologous chromosomes bearing linked genes mutually exchange corresponding parts Cryopreservation Freezing of cells or tissues in order to maintain viability with storage at very low temperatures Culture medium A substance or preparation used for the cultivation of living cells Cumulus oophoms A multilayered mass of follicular cells surrounding the oocyte. Cells of the cumulus are instrumental, via gap junctions, in nurturing the oocyte during growth and possibly in passing inhibiting factors necessary for deterring the resumption of meiosis. The innermost layer of cells is called the corona or coronal layer. This layer expands and presents a radiant pattern as oocytes mature in response to exogenous human chorionic gonadotropin (hCG) or a mid-cycle surge of luteinizing hormone (LH). Near ovulation, as they loosen and expand, cumulus cells are observed to retract from the zona pellucida of the oocyte, presumably cutting off the previously important cellularoocyte communication Cytokinesis A phase in mitosis or meiosis which involves the division of the cytoplasm Cytoplasmic fragmentation Disorganized fractioning of the cytoplasm, often in a manner that superficially resembles cleavage. Fragments may contain DNA, but they are more likely to contain no nuclear material at all Cytotrophoblast The thin inner layer of the trophoblast

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Desmosomes A type of junction that attaches one cell to its neighbor; one of a number of differentiated regions which occur, for example, where the cytoplasmic membranes of adjacent epithelial cells are closely apposed; consists of a circular region of each membrane together with associated intercellular microfilaments and an intercellular material Diandry Triploidy in which the extra haploid set is of paternal origin Differentiation The process of acquiring completely individual functions or characteristics, as occurs in the progressive diversification of cells and tissues of the developing embryo; the process by which a single pluripotent cell gives rise to a variety of different, more specialized cells Digyny Triploidy in which the extra haploid set is of maternal origin Dizygotic Pertaining to or derived from two separate zygotes, as in dizygotic (fraternal) twins DNA Abbreviation for deoxyribonucleic acid, a nucleic acid that is the carrier of genetic information for all organisms except the RNA viruses; DNA is found in all living cells Donor oocyte cycle A procedure of ART where a preembryo is formed from the oocyte of one woman (the donor) and transferred to another woman (the recipient) for purposes of establishing pregnancy Down’s syndrome (trisomy 21) A condition marked by an extra chromosome 21 or extra part of chromosome 21. Individuals are characterized by a small, anteroposteriorly flattened skull, short, flatbridged nose, epicanthal folds, short phalanges, widened space between the first and second digits of hands and feet, or variations in these stigmata, including moderate to severe mental retardation Ectoderm The outermost of the three primary germ layers of the embryo; cells of this layer will go on to form the epidermis and epidermal tissues (such as hair, nails, glands of the skin), external sense organs (ear, eye), some mucous membranes and nervous system of the adult Ectopic pregnancy A pregnancy in which the fertilized oocyte implants in a location outside the uterus, usually in the Fallopian tube, the ovary or the abdominal cavity Egg A term best reserved for a nutritive object frequently seen on the breakfast table; in humans, an oocyte Embryo The stage of the organism after development of the primitive streak; persists until major organs are developed. Once the neural groove and the first somites are present, the embryo is considered formed. In the human, the embryonic stage begins at

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approximately 14 days after fertilization and encompasses the period when organs and organ systems are coming into existence Embryogenesis The developmental process by which the embryo is formed

Embryologist Scientist with training and skills in handling spermatozoa, oocytes and preembryos in the laboratory; a scientist with specialized training in embryology Embryonic disc A structure in the stage of embryo development marked by the formation of ectoderm, mesoderm and endoderm Endocrinologist A medical doctor or scientist who specializes in disorders of the endocrine glands and the study of hormones Endoderm The innermost of the three primary germ layers of the embryo; cells of this layer will go on to form the digestive tract, epithelium of the pharynx, respiratory tract (except nose), bladder, urethra, lungs, etc. of the adult Endometrium The inner mucous membrane of the uterus, the thickness and structure of which vary with the phase of the menstrual cycle. It is functionally divisible into three layers: the stratum basale, stratum spongiosum and stratum compactum, the last two layers together forming the stratum functionale. The implanting human conceptus invades the endometrium Endoplasmic reticulum A cell organelle consisting of a complicated network of fine, branching and anastomosing tubules or spaces (cisternae) or isolated vesicles present in the cytoplasm of most cells. Endoplasmic reticulum forms a structural framework for the cells and a circulation pathway between the plasma membrane and nuclear membrane. Its surface may bear ribosomes, the site of protein synthesis Epididymis The elongated cordlike structure along the posterior border of the testis, whose elongated coiled duct provides for storage, transit, and maturation of spermatozoa and is continuous with the ductus deferens. It consists of a head (caput epididymis), body (corpus epididymis), and tail (cauda epididymis). Estrogen A female sex hormone responsible for secondary sex characteristics, the menstrual cycle and pregnancy Fallopian tube The oviduct; the tube leading from the ovary that is responsible for transporting the oocyte or fertilized oocyte to the uterus Fertilization The union of male and female gametes leading to the formation of a unique zygote

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Fetus The developing conceptus after the embryonic stage; the fetal period begins at the end of the eighth postovulatory week when greater than 90% of the more than 4500 named structures of the adult body have appeared. The fetal period persists until birth Follicle A structure in the ovaries that contains the oocyte and other cells

Follicle stimulating hormone FSH; a hormone released by the pituitary gland that stimulates the growth of the follicle and spermatogenesis Gamete The oocyte or the spermatozoon; a mature haploid reproductive cell; any cell which, upon union with another cell, results in the development of a new individual Gametogenesis The developmental process by which gametes are formed. In females the process is known more specifically as oogenesis; in males it is known as spermatogenesis Gap junctions Connections between cells which allow passage of small molecules and electric current. Gap junctions were first described anatomically as regions of close apposition between cells with a narrow (1–2 nm) gap between cell membranes. The variety in the properties of gap junctions is reflected in the number of connexins, the family of proteins which form the junctions Gene The functional unit of heredity which is a segment of DNA located at a specific site on a chromosome. A gene directs the formation of an enzyme or other protein Gene expression The full use of the information in a gene via transcription and translation leading to production of a protein and hence the appearance of the phenotype determined by that gene. Gene expression is assumed to be controlled at various points in the sequence leading to protein synthesis and this control is thought to be the major determinant of cellular differentiation in eukaryotes Genetics The science dealing with the passing of physical and chemical characteristics from parents to offspring and the impact of the environment on genes and genetic expression Genome activation (preembryonic) Transition of preembryonic development from control by maternally coded and stored messenger RNA from the oocyte to newly formed products of the preembryonic genome, including the paternally derived component. Genomic activation occurs at the 4–8-cell stage in human preembryos Germ cell In the male the testicular cell that divides ultimately to produce the spermatozoon; in the woman the ovarian cell that divides ultimately to form the oocyte

Glossary of terms

356

Gestation The period of time from conception to birth Gestational carrier A woman who enters into a social arrangement where she will carry a child created from another couple’s gametes with the expectation of relinquishing the child at birth (see Surrogacy for a similar arrangement involving carrying and giving birth to a genetically related child) Gestational sac A fluid-filled structure that develops within the uterus early in pregnancy. In a normal pregnancy, a gestational sac contains a developing embryo and, subsequently, the fetus Golgi apparatus, Golgi complex, Golgisystem A cytoplasmic organelle especially well developed in neurons and secretory cells, thought to play a role in the process of secretion. Golgi bodies can be considered the final packaging location for proteins and lipids. Each Golgi body consists of flattened membrane sacs. It is within the flattened membrane sacs that enzymes ready these proteins and lipids for shipment to specific locations. Vesicles form at the final region of a Golgi body when parts of the membrane begin to bulge. These vesicles then break away, via exocytosis, for the transport of these proteins and lipids to their final destination Gonadotropin Any hormone having a stimulating effect on the gonads. FSH and LH are two such hormones secreted by the anterior pituitary Gonadotropin-releasing factor A substance causing the pituitary gland to release LH and FSH Granulosa cells; membrana granulosacells Cells of the membrana granulosa lining the vesicular ovarian follicle which become luteal cells after ovulation; the layer of small cells that forms the wall of an ovarian follicle Haploid Possessing half the diploid or somatic number of chromosomes Hatched The end result of the hatching process where the blastocyst is completely removed from the zona pellucida Hatching The process by which the expanded blastocyst breaches and escapes through the zona pellucida; hatching must occur before implantation is possible HOMP High-order multiple pregnancy; more than two fetal hearts by ultrasound Human chorionic gonadotropin hCG; a hormone produced by the chorionic villi of the implanted/implanting conceptus; triggers the release of estrogen and progesterone Human menopausal gonadotropin A fertility hormone (drug) which is administered to promote follicular growth

Glossary of terms

357

ICSI Intracytoplasmic sperm injection. A procedure performed in IVF laboratories where a single sperm is injected into the cytoplasm of an oocyte to assist the fertilization process Immature oocyte An oocyte with chromosomes at prophase I (PI), a germinal vesicle-bearing oocyte Implantation The process involving attachment of the human blastocyst to the luminal epithelium of the uterus, its penetration through, and embedding within, the endometrium; in humans, this occurs 6–8 days after fertilization Implantation rate The proportion of gestational sacs by ultrasound divided by the number of conceptuses transferred to the uterus; given as a percentage Inner cell mass The cluster of pluripotent cells located at one point on the inner surface of the trophectoderm; these cells will form the body of the embryo after implantation Intermediate oocyte An oocyte with chromosomes at metaphase I (MI), characterized by the absence of both a first polar body and a germinal vesicle Intrauterine transfer; transfer;replacement The transfer of conceptuses to the uterine cavity for the purposes of establishing pregnancy IVF In vitro fertilization. A procedure wherein oocytes are fertilized and cultured outside the body Klinefelter’s syndrome A syndrome typically marked by a karyotype of 47, XXY (or XXYY, XXXY or XXXXY); or varying forms of mosaicism involving 46, XX and 46, XY cell lines. Individuals with Klinefelter’s syndrome experience infertility and usually exhibit variable degrees of masculinization, small testes and, sometimes, gynecomastia Leydig cells Cells of the testes that produce the male hormone testosterone Live birth The delivery of one or more living babies Luteinization The process by which a post-ovulatory ovarian follicle transforms into a corpus luteum through vascularization, follicular cell hypertrophy and lipid accumulation Luteinizing hormone LH; a hormone produced by the pituitary gland Male factor infertility Any cause of infertility due to low sperm count, motility, morphology or sperm function that makes it difficult for a sperm to fertilize an oocyte under normal conditions Mature oocyte An oocyte with chromosomes at metaphase II (MII), characterized by the presence of a first polar body

Glossary of terms

358

Meiosis A specialized form of nuclear division in which there two successive nuclear divisions (meiosis I and II) without any chromosome replication between. Each division can be divided into four stages similar to those of mitosis (prophase, metaphase, anaphase and telophase). Meiosis reduces the starting number of 4n chromosomes in the parent cell to n in each of the four daughter cells. Each cell receives only one of each homologous chromosome pair, with the maternal and paternal chromosomes being distributed randomly between the cells. This is vital for the segregation of genes. During prophase of meiosis I (classically divided into stages: leptotene, zygotene, pachytene, diplotene and diakinesis), homologous chromosomes pair to form bivalents, thus allowing crossing over, the physical exchange of chromatid segments. This results in the recombination of genes. Meiosis occurs during the formation of gametes in animals, which are thus haploid; fertilization gives rise to a diploid oocyte Menopause The ‘change of life’ or time when menstruation ceases, usually between the ages of 45 and 55 years Menstrual cycle Follicular phase The 12–14-day preovulatory phase of a woman’s menstrual cycle during which time a follicle grows Periovulatory Around the time of ovulation Luteal phase Post-ovulatory phase of a woman’s menstrual cycle. The corpus luteum produces progesterone, which causes the uterine lining to thicken to support implantation Menstruation The cyclic (monthly) shedding of the uterine lining in response to stimulation from estrogen and progesterone MESA Microsurgical epididymal sperm aspiration; surgical harvest of spermatozoa from the epididymis Mesoderm The middle layer of the three germ layers; cells of this layer will go on to form blood, blood vessels, lymphatics and lymphoid organs, gonads, peritoneum, notochord, pleura, pericardium, heart, kidneys, muscle, bone, cartilage, connective tissue, etc. of the adult Metaphase I oocyte An oocyte with chromosomes at metaphase I of maturation, characterized by the absence of both a first polar body and a germinal vesicle. An oocyte at an intermediate stage of maturation Metaphase II oocyte An oocyte with chromosomes at metaphase II of maturation, characterized by the presence of a first polar body. A fully mature oocyte. A secondary oocyte Miscarriage The loss of the products of conception from the uterus after a clinical pregnancy is established, but before the fetus is viable; spontaneous abortion Mitochondria One of the minute, spherical rod-shaped or filamentous organelles present in all cells. They contain many enzymes of the Kreb’s citric acid cycle and the electron transport

Glossary of terms

359

systems, hence are of primary importance in the metabolic activities of cells; mitochondria are the principal sites for the generation of energy and the only organelles other than the nucleus to possess DNA (mtDNA) Mitosis Division of a cell in which the two daughter nuclei normally receive identical complements of the number of chromosomes characteristic of the somatic cells of the species. Mitosis, the process by which the body grows and replaces cells, is divided into four phases: Prophase Formation of paired chromosomes, disappearance of nuclear membrane, appearance of the achromatic spindle, formation of polar bodies Metaphase Arrangement of chromosomes in the equatorial plane of the central spindle to form the monaster. Chromosomes separate into exactly similar halves Anaphas The two groups of daughter chromosomes separate and move along the fibers of the central spindle, each towards one of the asters, forming the diaster Telophase The daughter chromosomes resolve themselves into a reticulum and the daughter nuclei are formed, the cytoplasm divides, forming two complete daughter cells The term mitosis is used interchangeably with cell division, but strictly speaking it refers to nuclear division, whereas cytokinesis refers to division of the cytoplasm Monosomy A condition characterized by one less than the normal diploid number of chromosomes (2n–1) Monozygotic Pertaining to or derived from one fertilized oocyte or zygote, as in identical twins Morula Generally, the 8–16-cell stage when compaction commences until blastocyst formation; the stage commonly observed between 72 and 96 h after insemination. Some authors believe that the term morula is historically inappropriate for mammals Mosaicism A condition in which a conceptus or individual possesses two separate and distinct chromosome lines Multinucleation A state where cells possess more than one nucleus. Multinucleated blastomeres contain more than a single nucleus and are associated with preembryos demonstrating a reduced implantation potential Natural cycle Menstrual cycle that is not controlled or stimulated by exogenous drugs Natural selection The Darwinian principle that individuals or organisms with characteristics best suited to survival in a particular environment become a greater proportion of their species within that environment with each generation Neural tube The structure that forms from ectoderm on the dorsal side of a vertebrate that has been induced to form neural tissue. The neural tube goes on to form the spinal chord and brain

Glossary of terms

360

Nondisjunction The failure of a pair of homologous chromosomes to separate during the reduction division Notochord A tube of cells joined in the embryo foreshadowing the spine Nucleolus A rounded refractile body present in the nucleus of most cells which is the site of the synthesis of ribosomal RNA; the nucleolus becomes enlarged during periods of synthesis and atrophied during quiescent periods Nucleus The spheroid mass, enclosed in a thin membrane, which is the center for the synthesis of specific cellular proteins and the transmission of heredity traits; contains a nucleolus or several nucleoli, a diffuse nucleoplasm and DNA Oct4 A transcription factor essential for the maintenance of germ cell totipotence; Oct4 prevents germ cells from differentiating Oligozoospermia Production of an ejaculate with few sperm Oocyte The female gamete from inception of the first meiotic division until fertilization. In oogenesis, a cell which develops from an oogonium Oogonium The cell that gives rise to the primary oocyte during oogenesis. Oogonia proliferate by mitotic division during early fetal life Oolemma Plasma membrane of an oocyte Ooplasm Cytoplasm of the oocyte Ooplasm/cytoplasm descriptions Degenerative Non-viable Fragmented Exhibiting extracytoplasmic fragments or blebs Gmnular Exhibiting dense granules of darkish color Mottled Exhibiting ‘orange-peel’ granularity; ‘spotted’ Vacuolated Exhibiting spaces or small cavities within the cytoplasm of an oocyte or blastomere; may be the result of an aberrant endocytosis caused by oolemma instability Ootid The prezyote; the pronuclear stage oocyte before entrance into syngamy; the stage at which pronuclei are visible Ovarian stimulation The use of drugs (oral or injected) to stimulate the ovaries to develop follicles and ooctyes Ovary The female gonad; either of the two sexual glands in which oocytes are formed Ovulation The periodic production and discharge of an oocyte by the ovary

Glossary of terms

361

Ovum A female gamete or germ cell; an oocyte. The term ovum, which has been used for such disparate structures as an oocyte and a 3-week embryo, has no scientific usefulness Parthenogenesis The activation and subsequent development of a oocyte without fertilization; may occur naturally or through artificial stimulation Penetrated oocyte An oocyte that has been penetrated by a spermatozoon; strictly, one in which gamete plasma membranes have become confluent. The stage before pronuclei are formed. Penetration of the oocyte usually occurs within 3 h of insemination Perivitelline space Space surrounding the vitellus (technically, yolk; in this sense, the oocyte); the space between the oocyte and the zona pellucida; subzonal space. This space may possess the first and second polar bodies or extracellular fragments PESA Percutaneous epdidymal sperm aspiration; harvest of spermatozoa from the epdidymis without open surgery PGD Preimplantation genetic diagnosis. The procedure involving the removal of a blastomere or blastomeres from the developing preembryo for purposes of genetic analysis Pituitary gland The hypophysis; a small, oval endocrine gland lodged at the base of the brain that affects other endocrine glands Placenta A temporary organ which exchanges nutrients and wastes between mother and fetus and produces hormones needed to maintain pregnancy Pluripotent Cells capable of forming most tissues, i.e. cells of the inner mass (ICM). Although cells of the ICM can form every type of cell found in the human body, they cannot form an organism because they are unable to give rise to the placenta and supporting tissues necessary for development in the human uterus. These ICM cells are therefore ‘pluripotent’. Pluripotent stem cells undergo further specialization into stem cells that are committed to give rise to cells that have a particular function. Examples of this include blood stem cells which give rise to red blood cells, white blood cells and platelets; and skin stem cells that give rise to the various types of skin cells. These more specialized stem cells are called multipotent Polar body First polar body The structure extruded into the perivitelline space at the end of telophase I. Human chromosomes are divided between the oocyte and the first polar body (23 chromosomes, 46 chromatids, 2n DNA in each), those in the oocyte being attached to spindle microtubules. For a while after its formation, the first polar body remains connected to the oocyte by the meiotic spindle, forming a cytoplasmic bridge. Chromosomes within the first polar body may remain clumped together, may undergo a second meiotic division or may scatter within the cytoplasm; generally a nucleus is not formed. The first polar body contains cortical granules because of its extrusion before sperm penetration and cortical granule release

Glossary of terms

362

Second polar body The structure extruded into the perivitelline space after sperm penetration; contains 23 single-stranded chromosomes (1n DNA) after telophase II and may be nucleated Polarized; polarity The state or condition of having poles or possessing parts or regions of opposite or contrasting effects; cells that are polarized organize themselves or migrate to given poles, i.e. polar versus mural regions or inner cell mass versus trophectoderm, etc. Polar trophectoderm; mural trophectoderm Cells making up the trophectoderm of a blastocyst which are either polar (nearest or coming into contact with the inner cell mass), or mural (opposite or making no contact with the inner cell mass) Polyploidy A condition in which a conceptus or individual possesses one or more sets of homologous chromosomes in excess of the normal diploid set, as in triploidy (3n), tetraploidy (4n), hexaploidy (6n) or octoploidy (8n) Polyspermy The condition that occurs when an oocyte is penetrated by more than one fertilizing spermatozoon Post-transfer observation period The 1–3 days following intrauterine transfer when preembryos not frozen or replaced are evaluated daily to determine their suitability for freezing; also called the ‘post ob’ period Preembryo The conceptus during early cleavage stages until development of the embryo. The preembryonic period ends at approximately 14 days after fertilization with development of the primitive streak Pregnancy Clinical A pregnancy documented by ultrasonic investigation that shows a gestational sac in the uterus and requires dilatation and curettage (D&C) if miscarried Preclinical A pregnancy does not develop to the clinical stage despite initial positive testing Prezygote The pronuclear oocyte. The stage of development before syngamy when the term zygote becomes appropriate. Some authors refer to this stage as an ootid. Prezygotes are commonly observed 6–20 h after insemination or injection Primary oocyte The oocyte formed in the ovary before birth. Primary oocytes begin the first meiotic division before birth, but completion of prophase does not occur until after puberty Primitive streak A thick, opaque groove in the ectoderm that accompanies the emergence of the mesoderm and notochord Primordia The forerunners of organs or other structures in the embryo Primordial follicle An ovarian structure consisting of an oocyte surrounded by a single layer of granulosa cells

Glossary of terms

363

Progesterone A hormone produced by the corpus luteum and placenta that readies the endometrium for implantation and breasts for lactation, and maintains pregnancy Pronuclei Structures formed during fertilization from sperm and oocyte chromatin. Normal pronuclei are approximately 30 µm in diameter. See Prezygote Prophase I oocyte An oocyte with chromosomes at prophase I of maturation, characterized by a germinal vesicle Reactive oxygen species (ROS) Highly reactive substrates that sometimes form as byproducts of metabolism and are thought to cause permanent irreparable damage to the organism (preembryo, embryo or body) Recombinant DNA DNA artificially drawn from one species, combined with DNA from the same or a different species, and transplanted to the original or another species Refractile body An aggregation of lipid material and dense granules in the oocyte of approximately 10 µm size; associated with poor fertilization Secondary oocyte The oocyte after completion of the first meiotic division and arrest at metaphase of the second meiotic division. Also called a mature oocyte or metaphase II oocyte, this is the stage commonly associated with ovulated specimens or those collected from mature follicles for IVF. The secondary oocyte is characterized by a first polar body and no nucleus Seeding During cryopreservation, the process of introducing an ice crystal to the freezing solution in a controlled manner to prevent supercooling Selective reduction; multifetal pregnancyreduction A procedure used to decrease the number of fetuses a woman carries in order to improve the chances that the remaining fetus(es) will develop into a healthy infant(s). Reductions that occur naturally are referred to as spontaneous reductions Seminiferous tubules The small tubes of the testicles that produce spermatozoa Sertoli cells Cells in the seminiferous tubules that nurture developing spermatozoa Sex ratio, liveborn An expression of the number of live-born males in a population to the number of liveborn females, usually stated as the number of males per 100 females Somites Somites form from the lateral mesoderm. They are the precursors to the vertebrae and their associated muscles Spermatids The cells formed after the second phase of meiotic division (meiosis II) of the secondary spermatocytes

Glossary of terms

364

Spermatogonium A primordial germ cell that gives rise to a primary spermatocyte Spermatozoon A mature germ cell, the specific output of the testes. The generative element of the semen which serves to fertilize the oocyte, it consists of a head, neck, midpiece, and tail Spermiogenesis The transformation of a spermatid into a mature spermatozoon SSEAs Stage-specific embryonic (surface) antigens; carbohydrates associated with cell surface glycolipids, glycoproteins and proteoglycans. These antigens may be used as markers of cell differentiation and show speciesspecific expression Stem cells (embryonic stem cells) Cells with the ability to divide for indefinite periods in culture and give rise to specialized cells; pluripotent cells generated from the inner cell mass Stillbirth The birth of an infant with no signs of life after 20 or more weeks of gestation Subnuclei Nuclear or pronuclear fragments containing scattered, membrane-bound chromatin Supercooling During cryopreservation, the process of cooling to well below the freezing point without extracellular ice formation; when solutions supercool, cells do not dehydrate appropriately Surrogacy A social arrangement where one woman carries a child created from her own oocyte and donated semen for another woman (anonymous semen donor), man (directed semen donor) or couple (husband semen donor) with the expectation of relinquishing it at birth (see Gestational carrier for a similar arrangement involving carrying and giving birth to a genetically unrelated child) Synapsis The coming together in pairs of homologous chromosomes during meiosis Syncytiotrophoblast The outer layer of the trophoblast that invades the endometrium during implantation Syngamy The active union of two gametes in fertilization to form a zygote; the process of reorganization and pairing of maternal and paternal chromosomes in the zygote after pronuclear membrane breakdown Teratozoospermia Production of an ejaculate with a high percentage of abnormal forms (abnormal shapes) TESE Testicular sperm extraction; microsurgical extraction of spermatozoa from the testis Testis (plural: testes) The male gonad; either of the pear-shaped glands normally situated in the scrotum Testosterone The principal steroid hormone produced in men, responsible for secondary sex characteristics

Glossary of terms

365

Tight junctions Cell-cell junctions that seal adjacent epithelial cells (or epithelial-like trophectoderm cells) together, preventing the passage of most dissolved molecules from one side of the epithelial sheet to the other Totipotent Having unlimited capability to produce any type of cell. Totipotent cells have the capability to turn into (to ‘specialize’ or ‘differentiate’ into) the tissues surrounding the developing embryo, the embryo itself, and all of the tissues and organs that are present in the developed organism. The fertilized oocyte is totipotent until, perhaps, the 8-cell stage, meaning that its potential is total Translocation, chromosomal Interchange of chromatin between two or more chromosomes Triploidy A condition in which a conceptus or individual possesses three times the haploid number of chromosomes (3n) Trisomy A condition characterized by having one more than the diploid set of chromosomes (2n+1). Common trisomies have been given the names of those individuals first describing them: trisomy 13 (Patau’s syndrome), trisomy 18 (Edward’s syndrome); trisomy 21 (Down’s syndrome) Trophectoderm An outer, single layer of differentiated, polarized, epithelial-like cells forming the blastocyst; these cells ultimately give rise to the placenta and extraembryonic tissues after implantation. Polar trophectoderm Region of the trophectoderm in mammals that makes direct contact with the inner cell mass Mural trophectoderm Region of the trophectoderm in mammals not making contact with the inner cell mass Turner’s syndrome A syndrome marked by a karyotype of 45, X or varying forms of mosaicism involving 46, XX and 46, XY cell lines. Individuals with Turner’s syndrome experience ovarian failure and usually exhibit short stature and other somatic stigmata as described by Henry Turner Ultrasound A technique used in ART for visualizing the follicles in the ovary, the gestational sac or the fetus Varicocele (male) A varicose condition of the veins of the pampiniform plexus, forming a swelling that appears bluish through the skin of the scrotum, and accompanied by a constant pulling, dragging, or dull pain Vegetal pole/animal pole In an oocyte, the vegetal pole is opposite the animal pole. In contrast, the animal pole represents the region from which the polar bodies arise Zona pellucida The covering that surrounds the oocyte; believed to be produced largely by the surrounding follicular cells. In the human, the oocyte measures about 115 µm and the

Glossary of terms

366

thickness of the zona measures between 12 and 20 µm. The zona pellucida is covered externally by the corona radiata, which is a loose investment of granulosa cells from the ovarian follicle Zona reaction A process usually occurring during fertilization in which the chemistry and composition of the zona pellucida change to render it impermeable by other spermatozoa ZP1, ZP2, ZP3 Acidic glycoproteins localized in the zona pellucida of mammals; ZP3 is believed to be the major spermatozoon-binding protein and inducer of the acrosome reaction Zygote The 1-cell stage after pronuclear membrane breakdown and before the first cleavage. This stage is characterized by maternal and paternal chromosomes assuming positions on the first cleavage spindle and, thus, lacks a nucleus. Commonly observed 18–24 h after insemination or injection

Abbreviations and symbols used for embryology documentation Abbreviations abn

abnormal

c-c

cell-cell contact (noted just before compaction, as blastomeres become closely apposed)

clp

clear areas at periphery of cells

cont

contracted

deg

degenerative

dk, dksh

dark, darkish

f

faint

fert

fertilization

frag

fragmented

fz

fractured or broken zona pellucida

lg, med, sm

large, medium, small

m

many

mnb

multinucleated blastomeres

oval

oval shape

POB

post-transfer observation period

pb

polar body

pn

pronucleus/pronuclei

pvd

perivitelline debris

pvs

perivitelline space

rb

refractile body

sl

slight or slightly

subnuc

subnuclei

thk, thn

thick, thin

υ

very

vacs

vacuoles

zf

zona-free

zp

zona pellucida

Abbreviations and symbols

368

Symbols ∆

irregular shape (relates to shape of oocyte or blastomere) even (equivalent) sized blastomeres (relates to size) approximately even (approximately equal) sized blastomeres uneven (not equivalent) sized blastomeres

÷

cleaving (in the process of dividing) compacting compacted large pvs zona artifact

ô

bilayered zona

ö

porous zona

ō

dark zona mottled cytoplasm granular cytoplasm

√

good morphology

√√

excellent morphology very poor morphology

References Dorland’s lllustrated Medical Dictionary, 26th edn. Philadelphia: WB Saunders, 1981 Moore KL. The Developing Human, 3rd edn. Philadelphia: WBS aunders, 1982 O’Rahilly R, Muller F. Developmental Stages in Human Embryos. Washington, DC: Carnegie Institution of Washington, Publication 637, 1987 Society for Assisted Reproductive Technology/ Centers for Disease Control and Prevention. Assisted Reproductive Success Rates (http://www.cdc.gov/) Stedman’s Medical Dictionary, 25th edn. Baltimore: Williams & Wilkins, 1990 Steen EB. Dictionary of Biology. Savage, MD: Barnes and Noble Books, 1971 Veeck LL. Atlas of the Human Oocyte and Early Conceptus. Baltimore: Williams & Wilkins, 1986

Index abembryonic pole 64 acrosome reaction 259 actin filaments in adherens junctions 72 activation of oocyte 259 adherens junctions 62, 259 schematic presentation of 72 adhesion molecules 259 age (maternal), blastocyst transfer success and 101 multiple gestation incidence and 151 allocation 83–98 hypotheses of 84 American civil law 253 amino acid profiling 101 amino acids, effect on blastocyst 57 functions during preimplantation development 42 in sequential culture media 48, 49 ammonium production and nutrient uptake 60 amnion 259 aneuploidy 259 in in vitro blastocysts 144 annexin V 186 anterior-posterior polar axis 91, 92 apoptosis 185–202, 259 blastocyst cell number control by 85 blebbing and 194 characterization of 185 chromatin condensation at 187 condensed cytoplasm and 188 condensed nuclei 187 DNA fragmentation and 187 evidence for in preimplantation embryos 186 execution of 186 in 6-day blastocyst 196 incidence of 189 main features of 193 membrane changes at 186 morphological changes at 188 morphological features of 185

nuclear fragmentation and 187 phagocytosis and 189 regulation of 186 role of 189 timing of 189 apposition 203 aspermia 259 assisted hatching 161, 259 example 169 asthenozoospermia 259 asynchrony 145 autologous cumulus cells, use in coculture techniques 100 autologous endometrial cells, use in coculture techniques 100 autologous granulosa cells, use in coculture techniques 100 azoospermia 260 basic fibroblast growth factor 214 bax localization 199 bcl-2 localization 199 biomedical research, cloning for 234 biopsy, 260 of trophectoderm 250 birth weight and blastocyst transfer 103 blastocoel 260 collapse 167 enlargement of 115 fluid accumulation in 64 formation of 63 schematic representation of 113 zona drilling and twinning in 102 blastocyst 260 abnormal development of 108 age at transfer 99 apoptosis in 185 as experimental model 231–250 cell number and stage of development 100 and viability 85 in vivo cf. in vitro 100 cell differentiation in 83–98

Index cf. preembryo for transfer 145 cryobiology of 173 cryopreservation of 173–183 day-5 transfer cf. day-3 141 derivation of human stem cell lines from 214 development after ICSI 106 evaluation by staining 234 evaluation of quality 85 evidence for apoptosis in 186 examples of healthy 127, 128, 129, 130, 132, 156, 157 examples of unhealthy 132 fully expanded 91 gene expression in 86 gene regulation in 86 glycolysis and development 59 grade 1 expansion 91 grading criteria for 118, 119 grading systems 104 hatching of 159–171 high quality 150 identification of healthiest and IVF 12 implantation rates for 100 in vitro 99–137 in vivo fertilized 99 individual grading parameters 105 moral status of 251–257 nutrient uptake by 57 quality of 139–157 quantitative study of 85 schematic representation of, 113 sequential media and viability 60 serial development of 122, 123, 124 success of transfer and maternal age 101 success rates of transfer 102 time line for optimal development 107, 121 total cell number in 85 transfer and birth weight 103 transfer and sex ratios 103 transfer cf. preembryo transfer 12 with two separately cavitating segments 116 zona pellucida thickness and stage of development 99 ‘blastocyst breathing’ 167 blastomere 260 compaction and 61 formation of 24 hatching of after biopsy 170 multinucleated 155

371 polarity of 63 size and preembryo quality grading 25 totipotency of 28 blastomeres, exclusion of vacuolating blastomeres from compaction 71 nutrient requirements of 45 blebs 165 at hatching site 160 cf. projections 166 formation at apoptosis 194 human 194 multiple after freeze-thaw 171 murine 194 time-lapse example of 194 blood-borne pathogens 49 bovine oviductal cells, use in coculture techniques, 100 cadherins 260 system in adherens junctions 72 calcium, decompaction with lack of 73 role in compaction 63 canon law, see under law carbohydrate, functions during preimplantation development 42 in sequential culture media 48, 49 metabolism during preimplantation development 42 preference changes during preimplantation development 42 role in preimplantation development 86 cardiomyocytes 216 caspase activity 200 cavitation 231, 260 abnormal 82 advanced 80 blastocyst with two separately cavitating segments 116 central 74 exocentric 74 final stage of 81 following decompaction 79 process of 63 role of glucose in 64 schematic presentation of 74 serial development of 125 start of 79 cell adhesion molecules 204

Index cell allocation 83–98, 260 hypotheses of 84 models of 93 cell differentiation 83–98 dye studies 84 hypotheses of 84 models of 93 cell junctions, see under junctions cell lines of embryonic stem cells 213 cell-cell contact, full 77 minimal 75 moderate 76 stage of compaction 70 centriole 260 centrosome assembly 20 chaotic mosaicism 144 chimera 249 chorion 260 chromatin condensation, 187, 194, 195 chromosomal abnormalities, and fertilization failure 23 blastocyst development and 144 in in vitro blastocysts 144 Klinefelter’s syndrome 23 role in fertilization failure 23 chromosomes, combination of parental at fertilization 20 role in fertilization 23 cibacron blue 44 cleavage 260 after fertilization 20 fixing of first plane and polarization 83 preembryos after first 38 rate as selection criterion for transfer 143 rates and fertilization success 25 rates of in human preembryo 24 time of as selection criterion for transfer 143 totipotency at 84 cleavage-driven hypothesis 84 schematic representation of 93 clomiphene citrate use in ovulation induction 15 cloning 233–234, 260 coculture 260 techniques for 100 colchicine 260 collapse, partial 167 re-expansion and 160, 168

372 colony-stimulating factor-1 205 compaction 260 at day 4 94 cell-cell contact, differentiation of inner cell mass and trophectoderm at 69 exclusion of vacuolating blastomeres from 71 factors affecting 61 final stage 78 full 77 in 8-cell preembryos 71 minimal 75 moderate 76 normal cf. abnormal 61, 68 of morula 61 partial/incomplete 73 serial development of 125 stage of 70 timing of 61 conceptus 260 contraction during cryopreservation 175 Cornell criteria for cavitation 81 Cornell grading program for blastocysts 104 Cornell preembryo grading scheme 27 corona radiata 261 coronal layer 17 corticle granule release 20 premature and failed fertilization 20 Crabtree effect 43 crossing-over 261 cryopreservation 173–183, 261 blastocyst and 173 blastocyst before and after 181 blastocyst degeneration after 177 blastocyst freezing protocol 179 blastocyst thawing protocol 179 blastocyst after thawing 180 blebbing after 171 children born after 176 contraction and expansion during 175 degeneration after thawing 180 effect of day of freezing 176 first use 173 freezing of the isolated inner cell mass 176 natural cycle strategy for transfer 179 pregnancy rates after 175 programmed cycle strategy for transfer 179 protocols for 175 slow freezing and thawing 175

Index successful blastocysts 182, 183 technique 174 vitrification 174 cryoprotectants 174 culture media 261 coculture techniques 100 composition of 48 for human stem cell lines 214 formulation of 41 protein supplements in human 160 sequential 48, 49 toxicity of glucose to embryo 43 cumulus oophorus 17, 33, 261 cytochalasin-B 160 cytochalasin-D 100 cytokines, role in implantation 205 cytokinesis 261 of cleaving human preembryo 24 cytoplasm, condensation of and apoptosis 188 extensions in embryonic stem cell lines 221 fragmentation 154, 261 as selection criterion for transfer 143 immature cf. mature 17 vacuolization and apoptosis 189 cytoplasmic fragments, grading schemes for preembryos and 25 in preembryos 24 cytoplasmic string between inner cell mass and trophectoderm 114 cytotrophoblast 203, 261 decompaction 78 calcium and magnesium in 73 slight 79 desmosomes 62, 63, 261 schematic presentation of 72 TEM of 73 development, 2-cell stage 38 4-cell stage 38 6- to 8-cell stage 39 abnormal 108 after enzymatic removal of zona pellucida 106 after implantation 107 after zona pellucida manipulation 107 apoptosis and 185

373 cell number and stage of 100 chromosomal abnormalities and 144 in male cf. female preembryos 103 murine rates 160 preimplantation 231, 242 time clock for 99 diandry 261 differential staining 234, 248 differentiation 83–98, 261 cell types formed by 216 hypotheses of 84 of embryonic stem cells 215, 225–228 digyny 21, 261 causes of 21 diploid-aneuploid mosaicism 144 DMSO use in cryopreservation 174 dorsal-ventral polar axis 91, 92 dye studies of cell differentiation 84 ectoderm 262 EDTA, effect on blastocyst development 56 effect on embryo metabolism 43, 45 eight-cell stage of development 39 embryo 262 culture of human 47 physiology pre- and post-compaction 41 significance of glucose 44 significance of lactate 45 significance of pyruvate 45 toxicity of glucose to 43 viability assessment criteria and metabolism 46 embryo perfusion culture system 60 embryo twinning 233 by blastomere bisection 248 by blastomere separation 247 embryogenesis 203, 262 timetable of 210 embryoid body 225, 226, 227 embryonic germ cells 235 embryonic stem cells 213–229 cell types formed by 216 haracteristics of 213, 220, 221, 222 culture media for 214 derivation of cell lines 218 derivation of human stem cell lines 213 differentiation of 215, 225–228 murine cf. primate 214 Oct4 and 232 pluripotency of 235

Index typical morphology of 222, 223, 224 embryonic teratocarcinoma 214 embryo (2-cell stage) adenosine nucleotide ratio in 56 glycolytic activity in 56 pyruvate oxidation in 56 endoderm 262 endometrium 262 implantation and 203 ultrastructure of 204 endothelial cells 216 energy metabolism preimplantation 42 enzyme reactions 58 epidermal growth factor 205 expansion, degree of as a grading criterion 118, 119 during cryopreservation 175 experimental models of blastocysts 231–250 extended culture, day-5 transfer and 140 management of 140 extracellular matrix 205 fertilization, before oocyte reaches metaphase II 16 failure and chromosomes 23 importance of nuclear maturation in 17 polyspermic 21 preembryo cleavage rate and success 25 process of 19 pronucleus number and normality 22 fibroblast growth factor-4 84 flare protocols 16 follicle size and oocyte maturity 18 follicle-stimulating hormone use in ovulation induction, 15 four-cell stage of development 38 fragmentation 154 apoptosis and 187 as selection criterion for transfer 143 cytoplasmic 261 in preembryos 24, 25 DNA and apoptosis 187 in 3-day embryo 201 in 4-day embryo 197 nuclear and apoptosis 187 freezing, see under cryopreservation gametogenesis 263 gap junctions 62, 263 schematic presentation of 72

374 Gardner’s grading system for blastocysts 104 gene expression 263 changes in at apoptosis 188 in blastocyst formation 86 Oct4, 232–243 gene regulation in blastocyst formation 86 genome activation 263 glucose, amino acids and blastocyst uptake 57 during preimplantation development 42 effect on embryo 43 role in cavitation 64 significance of to embryo 44 toxicity of 43 uptake and viability, 59 glycolysis, activity in 2-cell embryos 56 blastocyst development and 59 in embryo 43, 44 rates as a marker for viability 101 viability assessment criteria and 47 glycoproteins in zona pellucida 159 gonadotropin, human chorionic, expression in cleavage stage preembryos 84 use in ovulation induction, 16 human menopausal, use in ovulation induction, 15 gonadotropin releasing hormone, long protocol cf. short protocol 16 pharmacological response to 15 use of agonists in ovulation induction 15 use of antagonists in ovulation induction 15 grading systems, criteria used for human blastocysts 118, 119 for blastocysts 104 for preembryos 25, 26 individual grading parameters for blastocysts 105 pronuclear grading system 152 selection criteria for day-3 transfer 142 granulation, after fertilization 22 at pronuclei stage 36 at prophase 135 granulosa cells 264 growth factors, role in implantation 205

Index

haploidy 145 hatching 99,159-171,264 after blastomere biopsy for preimplantation genetic diagnosis 170 after enzymatic zona removal 171 after manipulation of zona pellucida 170 assisted 161 beginning of process 165 blastocyst entrapment during 161 collapse and re-expansion and 168 enzymes and 159 failure of 171 heparin-binding epidermal growth factor and 160 initiation of 164 murine 159 rates of 160 premature 170 status of as a grading criterion 118, 119 trapping of inner cell mass at 115 hatching site of 160 helix-loop-helix transcription factor 86 heparin-binding epidermal growth factor 160 herniation after zona drilling 120, 121 herniation of trophectoderm 165 hoops 20 Hxt gene 86 ICSI 20, 264 blastocyst development after 106 hatching blastocyst after 168 oolemma piercing for oocyte activation 20 ooplasm disturbance for oocyte activation 20 preembryos after 153 trapping during hatching after 161 triploidy after 35 implantation 203-212 after delayed hatching 160 blastocysts, rates of 100 cell adhesion molecules and 204 cytokines and 205 development after 107 embryogenesis before 203 growth factors and 205 interactions involved in 209 successful blastocysts after cryopreservation 182, 183

375 window for 203 implantation rate 264 inner cell mass 264 abnormal 134 absent 133 cytoplasmic string with trophectoderm 114 derivation of cell lines from 218, 219 development of 83 differential labelling of 195 differentiation of 69 embryonic stem cells derived from 213229 formation of 94 fragmented nuclei in 196 freezing of the isolated 176 grading criteria for 118, 119 hatching in absence of 160 lack of visible 133 morphology of 95 optimal size and shape 105 pluripotency of 84, 99 position cf hatching site 164 proliferation rate cf. trophectoderm 85 rate of blastomere division and contribution to 100 schematic representation of 113 top-quality 120 trapping of during hatching 115 use in research 108 inside-outside hypothesis 84 schematic representation of 93 insulin growth factor binding protein-1 206 insulin-secreting cells 216 integrins 204 interleukins, role in implantation 205 Islamic tradition and the law 252 IVF, costs 11 ovulation induction for 15 twinning and 103 Jewish traditions and the law 252 junctions, formation of at cell-cell contact stage of compaction 70 formation of at compaction 62 schematic presentation of 72 see also under specific junction type Klinefelter’s syndrome 264

Index results of ART in 23 lactate, amino acids and blastocyst production 57 during preimplantation development 42 enzyme coupling and 58 significance of to embryo 45 uptake and viability 59 law, American civil 253 canon law and classical traditions 251 canon law and current traditions 252 Islamic tradition and 252 Jewish traditions and 252 natural reason and 253 use of blastocysts in research 253 legal status of blastocyst, see under law leptin 83 leukemia inhibitory factor 205, 214 LH use in ovulation induction 15 long protocols 16 MAG 205 magnesium, decompaction with lack of 73 role in compaction 63 maternal age, see under age matrix metalloproteinases (MMPs) 206 maturation score 18 maturation/meiosis/mitosis-promoting factor 246 maturity, assessment of 17 effect on fertilization success 18 fertilization success and 23 importance of 17 maturation score 18 metaphase I oocyte 19 metaphase II oocyte 18 of oocyte and role in digyny 21 prophase I oocyte 19 meiosis 265 membrana granulosa cells 17 metabolic control hypothesis 45 metabolism, impairment of 46 in male cf. female preembryos 103 nutrient requirements of blastomeres 45 of glucose in embryo 44 of lactate and pyruvate in embryo 45

376 requirements during preimplantation development 41–60 viability assessment criteria and 46 metaphase I, 265 maturation score description 18 oocytes at 34 metaphase II, 265 maturation score description 18 mature oocytes at 33 miscarriage and maternal age 101 mitosis 266 monosomy 266 monosomy 15 120 monosomy 21 144 monosomy 22 120 monosomy X 144 monozygotic twinning, blastocyst transfer and 101 blastocysts that led to 116, 117 rate of in IVF 103 time of 102 zona drilling and 102 morula 266 abnormal cavitation 82 advancing cavitation 80 assessment of for transfer 64 cavitation of 63, 94, 115 final stage of 81 start of 79 compaction of 61, 94 final stage of 78 decompaction of 78 description 61 extended culture techniques and transfer of 64 full compaction of 78 human 61–82 morphology of 64 multinucleation in 77 nuclei in 197 polar axes in 92 serial development of 122, 123, 124 murine 126 vacuolization, abnormal 82 normal 82 mosaicism 144, 266 Klinefelter’s syndrome 23 MUC-1 204 mucins 204 multinucleation 155, 266 as selection criterion for transfer 143

Index multinucleation in morula 77 multiple gestation, incidence 12, 140 maternal age and 27 odds of 151 reduction of incidence 139 risks involved 11, 103 see also twinning triplet rate 141 ultrasound of quadruplet pregnancy 150 ultrasound of triplet pregnancy 150 neural tube 266 neuronal cells 216 non-invasive amino acid profiling 101 nuclear maturation, importance of in fertilization 17 nuclear transfer 233 creation of fusion pair 245 cytoplast activation 246 embryo development 246 preparation of cytoplast 244 nucleolus, 266 alignment of 37 distribution of 152 as selection criterion for transfer 142 polarization of 83 nucleotide ratio in 2-cell stage embryo 56 nucleus 266 apoptotic 196, 197 condensation of and apoptosis 187 differential labelling of 195 fragmentation of and apoptosis 187 fragmented in inner cell mass 196 from 6-day blastocyst 196 in 4-day embryo 197 in 5-day morula 197 in 6-day blastocyst 195 numbers in 6-day blastocyst 198 visualization of in vivo 95 nutrient requirements for preimplantation development 41 nutrition, ammonium production and 60 composition of culture media 48 development and 47 requirements of blastomeres 45 uptake by blastocyst 57 Oct4 expression 86, 95, 267 in cleavage stage preembryos 84

377 in murine blastocysts 232 in primate blastocysts 232 in rhesus monkey 243 mutants for 232 oocyte 267 activation of for ICSI 20 aging, and late zona reaction 21 and spindle defects 21 assessment of maturity 17 at metaphase I 19, 34 at metaphase II 18, 34 at prophase I 19, 35 disorganized spindle in aged 34 fertilization before metaphase II 16 histological description 16 incubation of immature until metaphase II is reached 17 maturation score 18 nuclear maturation in 16 role of cumulus oophorus 17 sperm-penetrated 19 pronuclei in 36 oocyte donation 140 oogoniuim 267 oolemma 16, 267 fusion with spermatozoa 20 piercing of for oocyte activation 20 role in polyspermy prevention 21 ooplasm 16, 267 disturbance of for oocyte activation 20 ootid 267 definition 20 ovarian hyperstimulation syndrome 173 ovarian stimulation 267 ovary age and blastocyst transfer success 101 oviduct, nutrient levels in during preimplantation development 42 ovulation 267 ovulation induction protocols 15 oxygen consumption during preimplantation development 41 parthenogenesis 267 Ped gene 86 pentose phosphate pathway 44 perfusion culture system for embryos 60 perivitelline space 16, 267 phagocytosis 202 apoptosis and 189

Index sperm incorporation via 19 physiology of pre- and post-compaction embryo 41 pinopods 204, 212 plasminogen activator inhibitor 206 ploidy and zona drilling 120 pluripotency 268 of embryonic stem cells 215 of primate embryonic stem cells 235 polar axis, anterior-posterior 91, 92 dorsal-ventral 91, 92 polar body 268 extrusion after fertilization 22 first 33 pronucleus orientation and 152 role in fertilization 17 polarization, axes of 91, 92 establishment of 93 of nucleoli 83 role in cell differentiation 83 polarization hypothesis 84 schematic representation of 93 polyploidy 268 in in vitro blastocysts 144 polyspermy 268 after SUZI 21 block to 20 causes of 21 zona pellucida hardening and 161 post-compaction metabolic requirements 41 pre-compaction metabolic requirements 41 preembryo 268 2-cell stage of development 38 4-cell stage of development 38 6 to 8-cell stage of development 39 8-cell stage, compaction failure 68 compaction in 71 advancing cavitation 80 after first cleavage 38 after ICSI 153 cavitation of 63–64 cell number in vivo cf. in vitro 100 cf. blastocyst for transfer 145 cleaving cytokinesis of 24 morphology of 24 rates of cleavage 24 compaction of 61–63 cytoplasmic fragments in 24

378 decompaction in 78, 79 final stage of cavitation 81 full cell-cell contact in 77 good-quality that failed to reach blastocyst stage 155, 156 grading schemes for 25 incomplete compaction of 73 marker for viability 100 minimal cell-cell contact in 75 moderate cell-cell contact in 76 partial compaction of 73 proportion developing to blastocyst stage 99 quality of 24 selection for transfer 27 selection of 139–157 serial development of 122, 123, 124 serial photography on days 3–5 122, 123 start of cavitation 79 transfer cf. blastocyst transfer 12 viability of 12 pregnancy, incidence of loss 12 potential for and oocyte maturity 18 preembryo grading and success rates 26 rates after cryopreservation 175 risks of multiple 139 success rates and maternal age 101 preimplantation development 242 amino acid functions during 42 energy metabolism during 42 metabolic requirements during 41–60 nutrient requirements for 41 preimplantation embryo development gene 86 preimplantation genetic diagnosis 237–238 examples of healthy blastocysts after 132 hatching of blastomere after biopsy 170 herniating blastocysts after 121 importance of cell differentiation to 84 transfer delay and 64 prezygote 268 definition 20 progesterone and receptivity of uterus 107 PROH use in cryopreservation 174 projections cf. blebs and vesicles 166 pronase 161 hatching after use of 171 pronuclear grading system 152 pronuclear stage 20 pronucleus,

Index alignment as selection criterion for transfer 142 distribution as selection criterion for transfer 142 female 21,37 formation 20 grading system for 152 in sperm-penetrated oocytes 36 male 21,37 number of and normality of fertilization 22 orientation to polar body 152 period of growth 22 single 145 timing of formation after fertilization 22 prophase I 269 maturation score description 18 oocytes at 35 pyruvate, during preimplantation development 42 enzyme coupling and 58 oxidation in 2-cell embryos 56 significance of to embryo 45 quality, abnormal trophectoderm 136 blastocysts 139–157 development after ICSI and 106 evaluation of 85 with abnormal inner cell masses 134 with small inner cell masses 134 without visible inner cell masses 133 criteria used for human blastocysts 118, 119 day-3 morphology assessment of for transfer 144 degenerating trophectoderm 137 examples of healthy blastocysts 127, 128, 129, 130, 131 after preimplantation genetic diagnosis 132 examples of unhealthy blastocysts 132 grading schemes for preembryos 25, 26 high-quality blastocysts 150 improved identification with longer culture times 140 morphological grading scheme 26 nucleolus distribution and 152 of preembryo 24

379 poor 200, 201 top-quality inner cell masses 120 reactive oxygen species 269 research, use of blastocysts in 253 seeding 174, 269 selective embryo reduction 11, 269 sequential culture systems cf. coculture techniques 100 sequential media, blastocyst grown in 60 serial photography on days 3–5 of preembryos 122, 123, 124 sex, metabolism in male cf. female preembryos 103 sex ratios, after freezing 104 blastocyst transfer and 103 short protocols 16 six-cell stage of development 39 sodium levels in blastocoel fluid 64 sodium channels in trophectoderm cells 74 spermatozoa, effect on cumulus oophorus 17 fusion with oolemma 20 consequence of 20 immaturity of and fertilization success 23 incorporation 19 spindle, disorganized spindle in aged oocyte 34 normal meiotic 33 stage-specific embryonic antigen 214, 270 expression in human 87 expression in mouse 87 expression of SSEA-1 96 expression of SSEA-3 97 expression of SSEA-4 98 role in cell-cell interactions 86 role in compaction 63 staining, differential 234 stem cell research 108 strypsin 159 STST3 83 sucrose use in cryopreservation 174 sun-burst corona radiata 17 supercooling 174, 270 superoxide anion radical generation at hatching 159 SUZI, polyspermy after 21 syncytiotrophoblast 270 syngamy 36, 152, 270

Index

teratoma 228, 229 formation by embryonic stem cells 215 teratozoospermia 270 thawing, blastocysts after 180 degeneration after 180 protocols for 175 see also cryopreservation slow 175 technique 179 tight junctions 62, 270 in trophectoderm cells 74 schematic presentation of 72 TEM of 73 time line for blastocyst development 99, 121 for optimal blastocyst development 107, 108 tissue inhibitors of matrix metalloproteinases (TIMPs) 206 totipotency 28, 270 cf. polarity in mouse 83 loss of at compaction 61 loss of during cleavage 84 transfer, age of blastocysts at 99 blastocyst cf. preembryo 145 choice of optimal conceptus for 139–157 day-3, cf. day-5 141 history of 139 morphology of quality for 144 day-5, advantages of 140 history of 139 multiple gestation incidence and 140 success rate of 140 enzymatic removal of zona pellucida before 106 history of day-3 and day-5 139 monozygotic twinning and 101 multiple gestation incidence and number of preembryos transferred 151 natural cycle strategy after cryopreservation 179 programmed cycle strategy after cryopreservation 179 results in UK 142 results in USA 140, 141

380 selection criteria for day-3 142 success rates 102 of day-5 cf. day-3 140, 141 when to transfer single conceptus 145 transforming growth factor-β, 205, 206 expression 86 translocations 144 triplet rate 141 triploidy 271 after in vitro aging 35 cleavage rates in 25 incidence in ART cf. after natural intercourse 21 sperm concentration and 21 trisomy 271 in in vitro blastocysts 144 trisomy 13 120 trisomy 21 120,261 trophectoderm 271 abnormal 136 biopsy of 250 blebbing of 165 changes leading to hatching 159 cytoplasmic string with inner cell mass 114 degenerating 137 after thawing 180 development of 83, 114 differential labelling of 195 differentiation of 69 genes associated with 86 grading criteria for 118, 119 herniation of 165 poor quality 164 projections of 160, 166 proliferation rate cf. inner cell mass 85 role in pregnancy 99 schematic representation of 113 tight junctions between 73 trophoblast, invasion by 205 projections from 211 trophoblast cells 216 tumor-rejecting antigen-60 214 Turner’s syndrome 271 twin-twin transfusion syndrome 103 twinning, blastocysts that led to 116, 117 by blastomere bisection 248 by blastomere separation 247 cloning by 233 monozygotic, blastocyst transfer and 101

Index timing of 102 zona drilling and 102 two-cell stage of development 38 Tyrode’s solution 169 ultramicrofluorescence techniques 101 uterine lavage 99 uterus, nutrient levels in during preimplantation development 42 receptivity of 107 vacuolization, abnormal 82 cytoplasmic and apoptosis 189 normal 82 of blastomeres 71 viability and 135 Vero cells in coculture techniques 100 vesicles 165 at hatching site 160 cf. projections 166 viability, assessment of by metabolic criteria 46 glucose uptake and 59 grading systems for blastocysts 104 individual grading parameters for blastocysts 105 lactate uptake and 59 marker for 100 vacuolization and 135 vitellus 16 vitrification 174 zona drilling, effect on twinning 102 zona hardening 20 zona pellucida 16, 271 at metaphase I 34 blastocyst development after manipulation of 107 chemical alteration in to prevent polyspermy 20 drilling and ploidy anomalies 120 enzymatic removal of 106 hardening after sperm entry 161 hatching from 99 after manipulation 170 role in monozygotic twinning 103 structure and composition 159 thickness of and preembryo quality grading 25

381 and stage of blastocyst development 99 thinning of 115 zona reaction 20, 271 late in aged oocytes 21 ZP1 271 ZP2 271 ZP3 271